Insect Conservation: A Handbook of Approaches and Methods (Techniques in Ecology and Conservation)

Insect Conservation

Techniques in Ecology and Conservation Series Series Editor: William J. Sutherland Bird Ecology and Conservation: A Handbook of Techniques William J. Sutherland, Ian Newton, and Rhys E. Green Conservation Education and Outreach Techniques Susan K. Jacobson, Mallory D. McDuff, and Martha C. Monroe Forest Ecology and Conservation: A Handbook of Techniques Adrian C. Newton Habitat Management for Conservation: A Handbook of Techniques Malcolm Ausden Conservation and Sustainable Use: A Handbook of Techniques E.J. Milner-Gulland and J. Marcus Rowcliffe Invasive Species Management: A Handbook of Principles and Techniques Michael N. Clout and Peter A. Williams Amphibian Ecology and Conservation: A Handbook of Techniques C. Kenneth Dodd, Jr. Insect Conservation: Approaches and Methods Michael J. Samways, Melodie A. McGeoch and Tim R. New

Insect Conservation A Handbook of Approaches and Methods

Michael J. Samways Department of Conservation Ecology and Entomology, University of Stellenbosch, South Africa

Melodie A. McGeoch Cape Research Centre, South African National Parks

Tim R. New Department of Zoology, La Trobe University, Melbourne

1

3

Great Clarendon Street, Oxford OX2 6DP Oxford University Press is a department of the University of Oxford. It furthers the University’s objective of excellence in research, scholarship, and education by publishing worldwide in Oxford New York Auckland Cape Town Dar es Salaam Hong Kong Karachi Kuala Lumpur Madrid Melbourne Mexico City Nairobi New Delhi Shanghai Taipei Toronto With offices in Argentina Austria Brazil Chile Czech Republic France Greece Guatemala Hungary Italy Japan Poland Portugal Singapore South Korea Switzerland Thailand Turkey Ukraine Vietnam Oxford is a registered trade mark of Oxford University Press in the UK and in certain other countries Published in the United States by Oxford University Press Inc., New York © Michael J. Samways, Melodie A. McGeoch, and Tim R. New, 2010 The moral rights of the authors have been asserted Database right Oxford University Press (maker) First published 2010 All rights reserved. No part of this publication may be reproduced, stored in a retrieval system, or transmitted, in any form or by any means, without the prior permission in writing of Oxford University Press, or as expressly permitted by law, or under terms agreed with the appropriate reprographics rights organization. Enquiries concerning reproduction outside the scope of the above should be sent to the Rights Department, Oxford University Press, at the address above You must not circulate this book in any other binding or cover and you must impose the same condition on any acquirer British Library Cataloguing in Publication Data Data available Library of Congress Cataloging in Publication Data Data available Typeset by Newgen Imaging Systems (P) Ltd., Chennai, India Printed in Great Britain on acid-free paper by CPI Antony Rowe, Chippenham, Wiltshire ISBN 978–0–19–929823–5 (Hbk.) ISBN 978–0–19–929822–8 (Pbk.) 10 9 8 7 6 5 4 3 2 1

Acknowledgements We are enormously thankful to Ian Sherman and Helen Eaton of Oxford University Press for their encouragement and support. We also thank James Pryke and René Gaigher for their assistance with preparation of the figures. We are grateful too for the wonderful artwork of some of the equipment used in insect conservation drawn by Anni Coetzer. Adrian Armstrong, Mattias Larsson, Sharon Louw, Jürgen Ott, Nils Ryrholm, Glenn Svensson, and John Terblanche, individually or jointly very kindly supplied authored boxes on their specialist fields. Mark Robertson provided helpful comments on parts of the text. Many other researchers also supplied photographs and are acknowledged alongside their contribution. We wish you interest and enjoyment in your research, as we have had in ours over the years. Michael J. Samways Melodie A. Mcgeoch Tim R. New

This page intentionally left blank

Contents Introduction

xiii

1 Introducing insect conservation 1.1 1.2 1.3 1.4 1.5 1.6 1.7

The challenge: the relevance of sampling insects An historical perspective Ethics of insect conservation Ethics of collecting insects Ecological and evolutionary timescales The taxonomic challenge Summary

2 Taxonomy and curation of insects 2.1 2.2 2.3 2.4 2.5

Introduction: essential planning Selection of insect groups for study Taxonomic knowledge Taxonomic resolution Morphospecies and parataxonomists 2.5.1 The morphospecies approach 2.5.2 Parataxonomists 2.5.3 Voucher collections 2.6 Bioinformatics 2.6.1 Virtual voucher collections 2.6.2 Sample tracking 2.6.3 Taxonomy using barcoding 2.7 Specimen treatment 2.7.1 Sequence of steps for treating insect samples 2.7.1.1 Preservation 2.7.1.2 Mounting 2.7.1.3 Storage

2.8 Summary

1 1 2 3 6 10 13 15

17 17 19 21 25 27 27 29 30 31 33 33 34 35 38 38 46 53 57

3 Designing sampling protocols for insect conservation

59

3.1 Introduction 3.2 Defining the conservation question 3.2.1 Broad objectives in insect conservation 3.2.2 Refining the objective 3.2.3 Hypothesis formulation 3.3 Designing a sampling protocol 3.3.1 Sample grain

59 60 61 63 64 65 66

viii | Contents

3.4 3.5 3.6 3.7

3.3.2 Sample extent 3.3.3 Sample number 3.3.4 Sample independence 3.3.5 Spatial autocorrelation 3.3.6 Sample placement 3.3.7 Sampling coverage and intensity 3.3.8 Controls Pilot studies Coping with the unexpected Data management Summary

4 Collecting and recording insects 4.1 4.2 4.3 4.4 4.5 4.6 4.7 4.8 4.9 4.10 4.11

Introduction: What do you really need to collect? Named and nameless approaches to data portrayal High-canopy insects Low-canopy insects Grass/herbaceous layer Sampling flying insects Sampling insects on the ground Sampling insects in the leaf litter and soil Sampling aquatic insects Sound recording Summary

5 Measuring environmental variables 5.1 5.2 5.3 5.4 5.5 5.6 5.7

Introduction: selection of variables Conceptualizing environmental variable data Choosing the environmental variables Measuring air and soil environmental variables Measuring vegetation characteristics Aquatic environmental variables Summary

6 Estimating population size and condition 6.1 6.2 6.3 6.4 6.5

Introduction: the relevance of populations Assessing changes in population levels Significance of surveys Evaluating population structure Measuring population size 6.5.1 Some underlying principles 6.5.2 Temporal considerations 6.5.3 Spatial considerations and differences between sexes 6.5.4 Accuracy of estimates relative to feasibility

67 68 70 73 76 79 79 80 81 82 82

85 85 87 92 96 99 101 116 121 124 127 129

131 131 134 135 146 153 156 159

161 161 163 166 168 171 171 173 177 177

Contents | ix 6.5.5 Special considerations for threatened species 6.5.6 Direct counts 6.5.7 Transect counts 6.5.7.1 Line transects 6.5.7.2 Point transects 6.5.7.3 Belt transects

178 179 183 183 187 187

6.5.8 Quadrat counts 6.6 Populations and conservation status 6.6.1 Conservation status 6.6.2 Level of threat 6.7 Population distinctiveness 6.8 Population condition 6.9 Population modelling 6.10 Summary

188 189 189 191 202 204 205 206

7 The population and the landscape

209

7.1 7.2 7.3 7.4

Introduction: the relevance of landscapes Effects of landscape structure Resources and landscape geometry: the insect’s habitat Insect movements and MRR methods 7.4.1 MRR process 7.4.2 Capture 7.4.3 Marking 7.4.4 Release 7.4.5 Recapture 7.4.6 Outcomes from MRR studies 7.5 Landscape geometry and insect populations 7.6 Dispersal and dispersion (aggregation) 7.6.1 Activity patterns 7.6.2 Recording activity and abundance 7.7 Summary

209 211 217 218 226 228 229 232 232 233 235 240 243 244 245

8 Ex-situ conservation: captive rearing and reintroduction programmes

247

8.1 Introduction: conservation objectives 8.2 Farming or ranching insects for conservation 8.3 Process 8.3.1 Capture 8.3.2 Transport 8.3.3 Maintenance 8.3.4 Breeding 8.3.5 Release 8.4 Practical outcomes 8.5 Summary

247 252 253 257 258 259 260 264 268 271

x | Contents

9 Biodiversity and assemblage studies 9.1 9.2 9.3 9.4 9.5

9.6 9.7 9.8

9.9 9.10

Introduction: defining biodiversity in space and time The assemblage matrix The relational matrix Estimating sampling adequacy 9.4.1 Taxon sampling (accumulation and rarefaction) curves Species richness 9.5.1 Comparisons of species richness 9.5.2 Estimating species richness 9.5.3 Chosing a richness estimator Species abundance and density Species range and distribution 9.7.1 Biodiversity mapping Diversity indices 9.8.1 Species richness, evenness and dominance 9.8.2 Beta diversity 9.8.3 Functional diversity 9.8.4 Taxonomic diversity Phylogenetic comparative methods Summary

10 Studying insects in the changing environment 10.1 Introduction: drivers and methodology 10.2 Environmental change in context 10.3 Approaches to studying environmental change 10.3.1 Methodological approaches 10.3.2 Modelling 10.4 Bioindicators 10.4.1 Bioindication terminology and development 10.4.2 Taxa that make good bioindicators 10.4.3 Quantifying the indicator value of a species 10.4.4 Testing the proposed bioindicator and developing the bioindicator system 10.5 Biodiversity assessment 10.6 Biodiversity mapping 10.7 Summary

11 Key questions for insect conservation in an era of global change 11.1 Forms of environmental change 11.1.1 Habitat loss, fragmentation, and isolation 11.1.2 Disturbance, decline in habitat quality and the loss of critical resources 11.1.3 Impact of invasive alien species

273 273 275 278 278 280 282 282 282 283 284 287 288 292 292 295 298 300 302 303

305 305 305 306 306 307 310 311 312 318 321 322 325 331

333 333 333 335 336

Contents | xi 11.1.4 Climate change 11.1.5 Biotechnology, biological control, and sustainable harvesting 11.1.5.1 Biotechnology 11.1.5.2 Biological control 11.1.5.3 Sustainable harvesting

11.1.6 Aquatic and terrestrial systems contamination 11.2 Consequences of species loss for ecosystem function 11.2.1 Importance of diversity 11.2.2 Temporal perspectives, adverse synergisms and discontinuities 11.2.3 Species numbers, rarity and ecosystem function 11.2.4 The importance of food web connectance 11.2.5 Rare and threatened species as flagships 11.3 Lessons learned from diversity value: towards a synthetic management approach 11.4 Triage and restoration 11.5 Ecological monitoring 11.6 Engaging citizen scientists in insect conservation 11.7 Summary

Software References Glossary Index

337 346 346 351 352 353 354 354 354 355 357 358 359 364 365 368 371

373 375 405 425

This page intentionally left blank

Introduction Insects, as the most diverse group of animals with which we share our world, add substantial variety and diversity to the extraordinarily rare veneer of life on this biotic jewel we call Earth. Yet this insect diversity, the manifestation of millions of years of evolution, is under threat of attrition, as populations are lost, genetic diversity reduced, species go extinct, and the biomes and resources on which they depend succumb to human cupidity. It has been estimated that perhaps a quarter of all insect species are heading for extinction over the next few decades. In a mere blink of a geological eyelid, the ecological impact of humans is causing the demise of a vast amount of Earth’s biodiversity. This variety not only has intrinsic value but also may have much practical, utilitarian value which is only starting to be explored. Among the challenging issues in insect conservation is to establish the extent of diversity of the insect world, at genetic, population, and species levels. We then need to know which components of that diversity are being lost from which habitats and from where in the world. Insect diversity is so great and the challenge of its conservation so urgent, often we need to put in place actions that, through some value judgement, will conserve as much variety as possible rather than to treat each component species individually. To do this, we may need to use surrogates, which stand in for much of the diversity which we cannot adequately measure. Then, once we have put in place management actions that aim to conserve as many populations as possible, we need to establish how successful the conservation management actions have been. Insects, and many other terrestrial invertebrates, present some special challenges, simply because they are so varied, so diverse, and so unknown. They are also very much part of the fabric of terrestrial and freshwater ecosystems as we know them, and indeed are often intimately associated with plants. Insect ecology and conservation therefore goes hand in hand with plant ecology and conservation. This book aims to introduce approaches and techniques for insect ecology and conservation. While there are already some excellent introductory texts on sampling insects (although generally not with a specific conservation focus), the field of insect conservation has grown so much in recent years that there is a need for a contemporary and complementary text. Also, as there are some emerging techniques which are not in the traditional ecological techniques texts, we felt the need to produce a book that introduces students and researchers to approaches and techniques that can be used for both documenting and managing the conservation of insect variety. In general, this book is intended for the early-career researcher, with, say, a bachelor’s degree in biology, entomology or zoology who intends pursuing practical insect conservation work and writing a technical report, thesis, or research paper. We do not claim to be exhaustive, as new techniques, often

xiv | Introduction

Fig. 1 Layout of this book. The introductory chapters (2–5, 11) are closely interrelated and are the foundation for good planning of your project. The project may then be on a single species or on several species. Studies on the changing environment afftect both single species and species assemblages. Bold lines and arrows represent the most important linkages.

based on new ecological concepts, are continually emerging. The approach we take is summarized in Figure 1. Besides the ‘Introducing insect ecology and conservation’, there are four interrelated foundation chapters. The first of these, ‘Taxonomy and curation of insects’, emphasizes the importance of becoming familiar with your focal organisms, especially relative to the ecological and conservation questions being addressed, bearing in mind that ecology is a stepping stone to management for conservation. The next chapter, ‘Designing sampling protocols for insect ecology and conservation’, would seem, intuitively, to better sit after ‘Collecting and recording insects’. However, we wish to emphasize the crucial importance of planning the sampling before you go out to collect and record the subject insects. Then, as no insect lives in isolation, but rather is intimately interacting with its environment, it is essential to record appropriate environmental variables while sampling and recording insects (Chapter 5).

Introduction | xv

Your studies are then likely to be either on population aspects of single species, or, on the other hand, on the variety of insect species and their ecological relationships. Single-species studies are covered in Chapters 6–8. Assemblages (all species in one taxon, e.g. ants) and communities (interactions between species, e.g. food webs) are then dealt with in Chapters 9, 10, and 11. Of course, embedded in and sometimes even part of an assemblage/community study are aspects of population biology, hence the horizontal link in Figure 1. There is some overlap in the chapters to ensure consolidation of certain points and to strengthen linkages between chapters where it is crucial to do so. While we have made every effort to be fair and accurate, we cannot be held responsible for the effectiveness of the techniques, nor extensiveness of coverage, nor for individual actions relative to local, national, or international legislation.

This page intentionally left blank

1 Introducing insect conservation

1.1 The challenge: the relevance of sampling insects Insects are the most speciose class of animals, with nearly a million described species, and probably several times this number still to be discovered, diagnosed, and named, especially in the tropics (Grimaldi and Engel 2005). Their functional roles are enormous. They play major roles in virtually every terrestrial and freshwater ecosystem. The conservation of insects has major functional significance in maintaining the diversity of ecological processes on which all life ultimately depends. These roles are often difficult to quantify in economic terms. Nevertheless, in the USA alone, wild insects are estimated to be worth $57 billion per year by providing services such as controlling pests, pollinating flowers, burying dung, and providing nutrition for other wildlife (Losey and Vaughan 2006). The vast number of insect species, both described and undescribed, ensures their participation in an even greater number of food web interactions (Memmott 2000). Many of these interactions are highly specific, be they between herbivores and plants, predators and prey, or hosts and parasites. Our lack of detailed knowledge of the biology of most species renders their contribution to such integrated scenarios very difficult to evaluate. To assure continuing ecological integrity, we need to minimize losses. This approach is based on the precautionary principle, which means conserving as much as possible just in case there is any yet unforeseen value in genes, populations, species, or their interactions. To illustrate our lack of knowledge of the functional roles that insects play, in field samples we generally cannot even associate immature stages of insects with their corresponding adults, and these life stages are often functionally very different. The challenges of studying insects begin to multiply when we consider that many species can be sampled only during short periods of the year, and many disperse widely across the landscape and may not reside in the area in which they

2 | Introducing insect conservation

are sampled. Furthermore, many are small, cryptic, and intrinsically difficult to find and study. These challenges differ greatly from those facing investigators of vertebrates and plants. Past generations of entomologists have honed many of the traditional methods used to sample insects, largely through the transition of ‘collecting techniques’ to more quantitative or standardized methods, with many new approaches and refinements continuing to be developed (New 1998; Southwood and Henderson 2000; Wheater and Cook 2003; Leather 2005). Today, the purpose of sampling insects is not necessarily only for scientific advance. We now also sample insects for the purpose of ensuring we conserve as much insect diversity as possible (Samways 2005). We do this for insects in their own right, as well as to conserve the functionally important contribution that insects make to ecology and society. Let us now overview some facts, philosophy, and perspectives that enable us to pursue sampling insects with some broad and general insight.

1.2 An historical perspective The number of insect families has increased steadily over the last 400 million years (Mayhew 2007), with massive radiation during the early Carboniferous, just over 325 million years ago. The Permian extinctions took their toll on the terrestrial fauna, with loss of 8 out of 27 insect orders living at that time (Labandeira and Sepkoski 1993). Another major radiation event through the Triassic and Jurassic established the foundation for the huge insect diversity of today: 84% of the Tertiary insect families of 100 million years ago are still present today. Nevertheless, the meteorite impact that is thought to have caused the demise of the dinosaurs at the end of the Cretaceous, 65 million years ago, also affected the insects, with the implication that specialist feeders succumbed much more than generalists (Labandeira et al. 2002). Members of only about a third of the living insect orders eat plants. A few orders, such as Lepidoptera and Hemiptera, are predominantly herbivorous. To feed on plants in the first place was a huge evolutionary hurdle, simply because plant bulk is not nutritious and the surface of the plant is a hostile environment (Strong et al. 1984). Yet those insects that did adopt herbivory have scored handsomely, with today some 37% of all animal species being insects that feed on plants. There are strong suggestions that Cretaceous insects, both as herbivores and as pollinators, fostered an enormous feedback loop (co-evolution) where more plant species provided more opportunities for insect speciation. Other interactions, such as parasitism and predation, also became involved. The

Ethics of insect conservation | 3

resulting co-evolutionary process emphasizes that insect and plant ecology and conservation go hand in hand. Interestingly, there seems to have been little insect extinction during the Ice Ages, which were harsh and telling times for many large mammals and birds (Coope 1995). In response to the advance and retreat of the glaciers, insect distributions shifted southwards and northwards respectively (in the northern hemisphere) (Ponel et al. 2003). However, one result, which is critical for a conservation perspective, is that there was loss of genetic diversity along the northern margins of the northern hemisphere (Hewitt 2003). Mammals were important players on the landscape before the appearance of humans. Today we see the full effect of mammals only in a few places, such as the natural African savanna. However, shortly before tool-using humans began modifying and simplifying the landscape, there appear to have been large numbers of mammals creating a dynamic range of habitats. A resonance from these times is seen today where domestic livestock modify insect habitats, sometimes beneficially, but often not. Large mammal impacts on plants and on the physical environment, along with other factors such as fire, form an important ecological background for current insect conservation management. Indeed, for current conservation management a sense of the past is an important conceptual foundation (Figure 1.1). Hunting by tool-using humans led to dramatically changed interactions on the landscape. About 6000 years ago megafauna populations in many parts of the world plummeted and forests began to disappear. Although there is little definitive evidence for all but a few insect extinctions, some estimates can nevertheless be made. Mawdsley and Stork (1995) suggest that 11 200 insect species have gone extinct since the year 1600. The future also looks bleak, with McKinney (1999) pointing out that a quarter of all insect species are under threat of imminent extinction. Indeed, the losses may be greater for insects than for many other taxa, with insects in the UK, for example, declining more than vascular plants or birds (Thomas et al. 2004).

1.3 Ethics of insect conservation Studies of insects for conservation must begin with a firm set of values and clear objectives, not least because insects are often deliberately or unintentionally killed in the process. Furthermore, the basis for policy formulation of some conservation agencies is about the practical, utilitarian, or instrumental value of insects, with little consideration being given to the less tangible and higher values which constitute the ethical foundations of conservation. Yet, as outlined

4 | Introducing insect conservation Decades Millennia

Several millennia

Hundreds of thousand years

Tens of millions of years

Today

Glaciations Industrialization

Massive meteorite impact Supervolcanic (65 m.y.a.) cooling effects

First agriculture and deforestation

Fig. 1.1 It is conceptually valuable when undertaking insect ecology and conservation studies to have a sense of the past. In recent years, industrialization (along with intensive agriculture and urbanization) has made a major impact. By 6000 years ago, at least in Europe, deforestation and settled agriculture were well under way. Before that there were various dramatic natural events which were filters through which successful insect populations passed. These events give current species their particular biological characteristics and susceptibility to modern global environmental change.

by Haider and Jax (2007), incorporating a sound ethical base into conservation action can enhance its effectiveness. At the outset, we may ask why we should even bother with conservation of insect species, given that by far the majority of all species that have ever existed are today extinct from natural causes. Arguably, extinction is therefore the norm. There is one major difference between the situation today and yesterday: insects are currently becoming extinct faster than at any time in the past, largely because of the huge and sudden impact of humans, whose activities essentially diminish the capacity of even rapidly breeding organisms to adapt to more gradual changes in their environments. Thus it ethically behoves us to do something about slowing, and hopefully stopping, this extinction spasm. At the basal, utilitarian level of values, this means maintaining all the warp and weft of life which maintains ecosystems in a relatively stable, and more importantly, resilient state (Costanza and Mageau 2000) (Figure 1.2). In short, with no insects there can be no world as we know it.

Ethics of insect conservation | 5 Intrinsic value

Who cares about the earwig?

Utilitarian value Provisioning e.g. Food Water Fibre Fuel Regulating e.g. Eliminate climate regulation Water Disease Supporting e.g. Primary production Soil formation Cultural e.g. Spiritual Aesthetic Recreation Education

Fig. 1.2 The value of insects. Intrinsic value recognizes the importance of conservation for all organisms for their own sakes, irrespective of value to humans. Utilitarian value, on the other hand, recognizes the value of biodiversity to humans. Many of the categories of utilitarian value (based here on those of the Millennium Ecosystem Assessment 2005) are consumptive, i.e. of direct use, while some are non-consumptive, i.e. used but not consumed (such as many cultural values). The Precautionary Principle should be considered when dealing with insect species with consumptive value. In other words, care should be taken when using these taxa because any organism may play a role in maintaining a functioning ecosystem.

The utilitarian values of insects are varied. Some insects are important commodities, but insect values are not necessarily consumptive, as in the case of crop pollination and pest management. Protected areas were originally created largely for our aesthetic benefit. While they have ecotourism value, today they have crucial conservation value as well. Mostly, the subjects of nature reserves and parks are rare and charismatic species that readily foster interest and are tourist drawcards. This non-consumptive yet utilitarian value is also playing an increasingly important role in insect conservation (Hill and Twist 1998). These values, however, remain essentially human-orientated. An alternative value system is to consider the intrinsic value of organisms (Figure 1.2). This view sees insects and all their interactions as having the right to be, irrespective of their usefulness to us. We and they have entwined destinies, with rare and inconspicuous species also having a place on the conservation agenda. In turn, this approach raises the profile of insects as subjects worthy

6 | Introducing insect conservation

of conservation, as it also does with other cryptic organisms. However, insects are also particularly important components of many terrestrial ecosystems, and these ecosystems have systemic value. Conservation must therefore be directed towards protecting the natural biotic components and interactions in these ecosystems (Rolston 1994). Many conservation organizations are largely driven by utilitarian values. However, the Red List, which is the most authoritative list of threatened species, and produced by the World Conservation Union (IUCN), in particular has a very strong flavour of intrinsic value. A wasp on the Red List can, at least theoretically and based on information available, receive the same amount of attention as a whale. In other words, the Red List system encourages ‘equal value’. Of course, in practice, this is not always the case, simply because most people are more concerned for a blue whale than for a lesser-spotted fig wasp. With an increase in environmental awareness worldwide and a rise in collective human insight, the profile of insects is increasingly being raised in general conservation planning. Biodiversity conservation is now being more honestly brokered than in the past, as a wider range of organisms are being included, rather than mainly large and charismatic vertebrates.

1.4 Ethics of collecting insects Collecting is an all-embracing term for taking specimens from the wild. More formal sampling involves taking a subset of the whole, so as to have a meaningful representation of the whole. In practice, and ethically, this can be a difficult decision for the insect conservationist, as sampling commonly equates to killing, to enable identification, naming, and counting of specimens. It is therefore important to collect only enough individuals to answer the key question being posed by your investigation, and to minimize unnecessary deaths or harm to individuals. Furthermore, when very rare or threatened species are studied, the capture or removal of any individuals may be prohibited or a serious threat. General sampling at a site, for example to estimate species diversity, must be guided by possible harm to any threatened or susceptible taxa known to occur there. An additional consideration is that the sample must be manageable, there being no point in collecting so many individuals that they cannot be sorted and analysed. Intuitively obvious as this may seem, it is nevertheless surprising just how many studies in insect ecology and conservation flounder on this point. So, to restate our central theme here: it is critical to ask precisely what it is that you are researching? The answer to this question then determines your sampling protocol. How many insects you then sample also depends on the rarity of the

Ethics of collecting insects | 7

focal species and whether it is threatened or not. The insect conservationist thus engages in collection parsimony, where individuals are taken or disturbed only to a minimum to answer the probing question. A corollary is that you need to bear in mind that any physical sampling method is only one window on the world. Methods need to be chosen which address your research question and give a true reflection of the world you are investigating, while not harming your subject populations. Several general codes of conduct for insect collectors combine the ethical and practical considerations needed to ensure a responsible approach (Box 1.1 is an example). Although these codes were initially designed mainly for hobbyists, the major points included apply, sometimes even more particularly, to those engaged in scientific sampling. Box 1.1 Code for insect collecting With the increasing loss of habitats a point has been reached where a code for collecting should be considered in the interests of the conservation of insects. Whereas in the past collecting had only a trivial effect on insect numbers, today in some parts of the world, habitat loss has become so acute that collecting may have a critical extra impact on species. By subscribing to a Code for insect collecting, entomologists show themselves to be a concerned and responsible body of naturalists who have a positive contribution to make to the cause of conservation. All entomologists should in principle accept the following code, and try to observe it in practice. 1. Collecting—general 1.1 No more specimens than are strictly required for any purpose should be killed. 1.2 Readily identified insects should not be killed if the aim is to examine them for aberrations or any other purpose: insects should be very carefully handled alive and then released where they were captured. 1.3 The same species should not be removed in numbers year after year from the same locality. 1.4 Supposed or actual predators and parasitoids of insects should not be unnecessarily destroyed. 1.5 When collecting leaf-mines, galls, and seed heads do not collect all individuals that can be found; leave as many of these habitat units as possible for the population to recover. 1.6 Consideration should always be given to digital photography as an alternative to collecting (in some cases, such as with butterflies, this alternative may even be convenient).

8 | Introducing insect conservation 1.7 Collect specimens for exchange, or for sending to other collectors, sparingly or not at all. 1.8 Insects being used for commercial purposes should be either bred or obtained from old collections, and not collected from the wild. 2. Collecting—rare and threatened species 2.1 Rare species should be collected with the greatest of restraint. 2.2 Threatened species should generally not be collected at all, unless with very good reason and with all the appropriate collecting permits. 2.3 Collectors should look for new localities rather than collect local or rare species from well-known and perhaps overworked localities. 2.4 Previously unknown localities for rare and threatened species should be brought to the attention of local conservation authorities, so that species protection measures can be initiated. 3. Collecting—lights and light traps 3.1 The catch at or in a light trap should not be automatically killed. 3.2 Live trapping should be practised wherever possible, so that the trapped insects can be released back into the wild at the same habitat after examination. 3.3 Particular care should be given to catching very rare or threatened species. If this happens accidentally, traps should in future be relocated. 3.4 Light traps should be positioned so as not to annoy other people. 4. Collecting—permission and conditions 4.1 Always seek permission from the landowner or property occupant when collecting on private land. 4.2 Always comply with any conditions specified by the permit-granting agency. 4.3 When collecting (accompanied by an appropriate permit) in nature reserves, always supply the final species list to the relevant authority, as this information is often very valuable for management of the reserve. 5. Collecting—avoidance of damage to the habitat 5.1 Minimize disturbance to other wildlife as far as possible, especially to nesting birds and rare plants. 5.2 When beating bushes and trees for insects, beat the branch rather than thrash and damage the foliage. 5.3 When searching under bark and in leaf litter, always replace the material back as you found it. Always leave some bark or litter in the area, so some populations remain undisturbed. 5.4 Overturned logs and stones should always be returned to their original positions.

Ethics of collecting insects | 9 5.5 Aquatic plants, mosses, and liverworts should always be replaced in their original positions. 5.6 When sampling twigs, leaves, or inflorescences always use secateurs rather than simply breaking or tearing the plant part. 5.7 Be very selective in choosing sites for treacling, so as not to leave unsightly marks on trees. 5.8 Be particularly sensitive to and aware of any rare or threatened species in the area which could be disturbed by your activities. 6. Breeding 6.1 Breeding from a fertilized female or pairing in captivity is preferable to taking a series of specimens in the field. 6.2 Never collect more larvae or other livestock for rearing than can be supported by the available supply of food plant. 6.3 Unwanted insects that have been reared should be released in the original locality, not just anywhere. Alien insects should NEVER be released. 6.4 Do not attempt to establish a new colony or population in a new area without first consulting local conservation authorities. Based on the Invertebrate Link code (www.benhs.org.uk/code.html)

Collecting also depends on assessing the time needed to process the collected specimens, and facilities available for housing them (see Chapter 2 for more details). Most multi-species studies must be supported by a voucher collection. This is a range of specimens for immediate and regular use as a reference for yourself, and as reference for researchers in the future. The collected species in such studies may at first not be identified, and simply be recognizable morphological forms, called morphospecies (see Chapter 2 for more details). It is much easier to remember nicknames rather than numbers, so it is often useful to give your morphospecies in the voucher collection vivid, personalized names in lieu of the scientific names which can be substituted later. For most studies, these morphospecies will be named by taxonomic specialists. For some taxonomic groups, and where there are good keys, it may be possible for you to do some of the identifications yourself. Nevertheless, where precise scientific identifications are required, your conclusions should be verified by a specialist. Designated morphospecies may, in fact, turn out to be two or more true species, one or more morphological or ecological forms, different life stages, or different sexes. If this happens, it does of course call into question the validity of analyses conducted on morphospecies. Finally, the voucher collection should be kept safe and curated as a formal museum collection in perpetuity (see Chapter 2 for more details). Sometimes it is also essential to undertake contingency collecting. This means collecting for future potential projects. The prime example is for barcoding and other

10 | Introducing insect conservation

molecular studies. When this is the case, at least five individuals (or in the case of rare and threatened species, a single leg each from five different individuals from the same local population) should be collected and kept in absolute alcohol or acetone. Other potential uses include image analysis (for fluctuating asymmetry studies, for example) or for a pictorial database. For traditional morphological studies, ideally you should retain about 12 specimens (to illustrate variation about the mean) although for pictorial purposes only 2–3 specimens in good condition would be required. Before any sampling takes place, it is essential to investigate the legal requirements for collecting permits for the places (e.g. national park) or taxa (e.g. protected species) you intend to sample and, if necessary, to obtain these. There are now many cases where insect collectors, including bona fide researchers, have inadvertently—or even deliberately in some instances—transgressed the local legislation regarding procurement of specimens. Always ask the local conservation authority about what is required before you start collecting or sampling, and allow plenty of time for any necessary permit applications to be prepared and processed, as this can take up to several months. One further consideration is that you may wish to send specimens across international borders in which case you may need a CITES (Convention on International Trade in Endangered Species) permit. Again, start by asking your local conservation officer for advice on any import or export permits needed.

1.5 Ecological and evolutionary timescales Sampling is a snapshot, in both time and space. Insects often have short generation times and their evolution can be very rapid (Mavárez et al. 2006). A conceptually good starting point is that short term (seconds to months) can be considered as ecological time and longer term (usually many tens of years) as evolutionary time. Over time ecological processes gradually translate into evolutionary processes. This distinction is conceptually important. Whereas ecological studies are essentially short-term, conservation must consider not only the current ecological situation, but also long-term evolutionary potential. Conservation is not just about today, but about the future as well. However, one must be very careful not to put absolute time frames on ecological vs evolutionary timescales, as not only do they blend into each other, but more and more findings are showing that evolution, especially among insects, can happen very quickly indeed. Thus we are dealing with a conceptual spectrum, and not a finite timescale. Because sampling is such a brief episode in the complex ever-changing milieu of life, it is critical to be crystal clear on your goals. First, is it a spatial study, a temporal study, or both that you are planning (Figure 1.3)? Spatial sampling

Ecological and evolutionary timescales | 11 Global 10 000 km

Climate Soil moisture variations Seasonal vegetation cycles

1000 km

100 km

10 km

Spatial scale

1 km Local Second

Origin of earth and life

Carbon dioxide variations

Atmospheric convection

Glacial periods Speciation Extinction events Soil development

Nutrient cycles

Atmospheric turbulence Minute

Day

Year

Century

10 tya

1 mya

1 bya

Global Climate change

Habitat alteration and destruction, pollutants

Local

Single event, point-source disturbance Minute

Century Temporal scale

Fig. 1.3 The space–time scales of Earth system processes (above) and the scales at which studies of human-induced environmental change impact on insect biodiversity are conducted. The shaded area is the space–time scaling zone within which investigations are likely to have maximal predictive power. When starting an insect conservation research project, you need to be very clear on the aims or goal of the study. As part of the planning, you need to have a clear vision of the spatial scale of the study (e.g. small scale: comparison of insects on individual trees; large scale: comparison of aquatic insects across ponds in a region, even perhaps a global study). The spatial scale also depends on resources available, physical and financial, relative to time available. Your study may only be spatial, but ideally samples should be taken over time, at least to get some indication of seasonal changes. True temporal studies are conducted over many years. Seasonal studies and those conducted over a few years take place in ecological time. Studies examining, for example, thousands of years of subfossils of beetle elytra in soil strata, are considering changes over longer, evolutionary time (redrawn and modified from McGeoch 1998).

12 | Introducing insect conservation

takes place at various scales from local to global, making it necessary to interrogate the scale of your investigation. Are you, for example, comparing insect assemblages on different trees in the same local area, or are you comparing insect assemblages on particular trees but at different locations? It may even be a larger-scale study, which then calls for careful consideration of the practicality of simply getting to the sites. Whatever the scale, the study will require adequate replication, often at more than a single scale (see Chapter 3). Investigating temporal changes has its own set of challenges (see Chapter 3). Nature, virtually by definition, is always changing. There are nested changes, from diurnal (24 hours) to seasonal, then to roughly decadal (relative to sunspot cycles) and to longer term still. Overlying these regular changes are stochastic ones, where conditions may become unusually dry or wet, or hot or cold. Such out-of-the-blue impacts can be major for insects, whether directly, such as by affecting survival or behaviour, or indirectly, by affecting their mutualists, predators, parasitoids, and pathogens. Time of day can be very critical for many species of insects, which often have strict diurnal behavioural patterns. Such behaviour may change when conditions are particularly adverse. Dragonflies, for example, may be on the wing at midday on moderately warm days, yet retreat to shade at the same time of day on very hot days. Closely related species may exhibit very different patterns of diurnal activity. Long-term changes may be easy to track but may be difficult to interpret, because the environmental variables that affect insect populations are interrelated (synergistic) (Figure 1.4) and change in intensity. For example, as rainfall patterns change often so does temperature, and these in turn may interact with human pressures such as intensity of agricultural practice. Such a complex of synergistic factors is believed, for example, to be driving changes in butterfly populations in Europe (Warren et al. 2001). Despite all these causes of variation, a temporal study over a year can often provide valuable initial insight. However, such a study can be only a first approximation of seasonal changes in the focal insect population. Nevertheless, it can be very useful for focusing more detailed studies, such as measurements of population density or movement patterns. But rarely, if ever, is a single season or year of study enough to provide definitive information on which sound management plans can be based. In summary, even studies at small spatial scales present logistical and analytical challenges. As the spatial scale gets larger, practicalities, such as simply travelling to sites on time, become a major consideration. Temporal scaling also has its set of challenges. Genuinely long-term studies need team work, although smaller, short-term studies carried out by one person, say, over three days, at different times of the year (e.g. each month), can provide real insight into seasonal

The taxonomic challenge | 13

Fig. 1.4 Many impacts on insect populations, assemblages, and communities are multiple and synergistic, and change over time. These synergistic impacts need to be thought through very carefully to ensure that all environmental variables are accounted for in your study. Pictured here is the Basking malachite damselfly Chlorolestes apricans, a highly threatened species of the Eastern Cape, South Africa, at one of its last remaining intact habitats with an abundance of natural, low-growing riparian vegetation. Elsewhere, the species is suffering from the key threat of invasive alien trees, which continue to grow and spread along the river bank, shading out its sunny perches. The presence of the trees, in turn, leads to other adverse impacts, including shading out of the indigenous plants which are its oviposition site. The alien trees also encourage cattle to shelter under them, so breaking the stream banks and silting the water.

changes of insect populations and assemblages, essential for their conservation. Long-term studies are inherently difficult to undertake, not least because most granting agencies operate on a funding cycle of only a small number of years. A final consideration when planning a study is that future generations may find your baseline results very valuable for comparison. This is why it is essential when writing up to record and save information on all aspects of your study in detail (exact location, sampling methods, species recorded, etc.) so that such direct future comparisons can be made.

1.6 The taxonomic challenge The world of insects is so vast and complex that many assemblage- or communitylevel studies, particularly in warmer countries, are dealing with species which we currently cannot name. Yet we still aim to understand and conserve this nameless

14 | Introducing insect conservation

mass. This is the taxonomic challenge. Dealing with it involves focusing the study, both spatially and temporally. It also means being clear on the exact objectives and outcomes of the study. In practice, this usually involves doing some preliminary sampling before embarking on the full project. Although armchair planning may be the starting point, it is only when you get into the field and start some of the actual sampling that it becomes possible to assess whether the practical methodology is feasible and suitable for achieving your objectives (see Chapter 3.4). Once you have done such a pilot study, only then can you revise your objectives and replan. This may mean changing what you set out to do originally, which is far better than continuing uncritically and ending up with a pile of data of little value. A

A

B

C

A

B

C

D

Species

Genus/family

Order

Recognized, but no scientific name = morphospecies Recognized, and scientifically identifiable = species

Fig. 1.5 The morphospecies concept. When sorting insect samples, it is best to start by sorting specimens into higher taxa, such as order or family/genus. Simply using these higher taxa rarely has any significant ecological value because such higher taxon groups do not provide sufficiently high resolution information for conservation. There is much more information in samples if you can identify specimens to lower taxonomic levels, particularly to species level. However, many species are undescribed and many others cannot be identified and named without consulting a taxonomic expert. Non-experts can sort (represented by the two-pointed arrow) the specimens into putative species (termed morphospecies) and give them an interim or working name, e.g. Coleoptera, Carabidae sp. 1. This is the morphospecies approach. Here, recognizable species A is sorted, with other specimens (B and C) into morphospecies (upper arrowhead). Where there is a dearth of taxonomic knowledge or expertise, they will remain as morphospecies. However, it may be possible to name the three species and assign them to known scientific species (lower arrowhead). Both these outcomes enable analysis of the ecological data, although species names clearly have much more taxonomic and ecological value than the casual names of morphospecies.

Summary | 15

Critical to this feasibility study is the dimension of your taxonomic challenge. It may only be possible to sort samples to a higher taxon (e.g. family), labelling the component morphospecies (specimens recognizable as morphological entities and thought as possibly true species; see Chapter 2.5.1 for more details) of that higher taxon as sp. 1, sp. 2, . . ., sp. n. Such a study has two weaknesses. First, among insects, higher taxa are generally poor surrogates for species patterns and responses to change. Second, when doing assessments for whatever reason, the use of specimens formally identified to species level makes results repeatable. This is not possible with morphospecies (but see Chapter 2 for some solutions). Using higher taxa also has reduced value for conservation, especially of endemic species. Again, this is because without species names, it is not possible to compare samples from one study to another and so see how the status of species is changing relative to environmental change. Nevertheless, morphospecies can be excellent when undertaking a comparative, one-off study of, let’s say, contrasting landscape patches and how the profile of the insect assemblages is changing. Indeed, in geographical regions where the insect fauna is poorly known, use of morphospecies may be the only reasonable option. Another possibility is to use a mixture of actual named species and morphospecies. This at least provides some taxonomic resolution yet also enables the project to continue without being held up by taxonomic challenges.

1.7 Summary Insect diversity has increased over geological time, with insects today being more speciose than any time in the past. Their success has been associated with that of plants, meaning that insect conservation goes hand in hand with that of plants. Insect extinctions are becoming more widespread, making it important that we do something now to mitigate this trend. At the outset of a study, it is important that we are aware of the ethical foundation for why we are doing it. On the one hand, motivation for insect conservation can be utilitarian (we conserve because it is useful for us to do so), or, on the other hand, we conserve because of higher values which respect insect survival in its own right. This translates into whether we are conserving, say, an attractive butterfly species per se, or whether the reserve area for that butterfly is also significant for a whole range of other insect species which may not be glamorous or even particularly visible. We sample insects to get a meaningful picture of the whole without actually measuring that whole. We choose a subset to provide ecological information and to enable us to make conservation decisions. As insects and their world are so complex, it is essential when starting a project to be absolutely clear on the

16 | Introducing insect conservation

exact research and conservation goal. We also need to be very sure about exactly how much time is available to achieve that goal. Sampling thus has to be done bearing the end in mind, so that we sample effectively and leave enough time for analysis and writing up. Collection of data and specimens is therefore done parsimoniously, but may also involve collecting for unforeseen eventualities, such as for future DNA studies. Although insect ecological studies are largely about the present, insect conservation also considers the future and evolutionary possibilities for insects. As sampling needs to be tuned to the research question, it means being clear on the spatial and temporal scales of our study from the outset. We also need to have realistic expectations and ensure that our time and funding is sufficient to undertake the planned work. This means developing an initial plan but then going out into the field and doing some preliminary sampling to see what is actually feasible. Insect ecology and conservation studies inevitably have major taxonomic challenges (see Chapter 2). Higher taxon surrogates, such as orders or families, rarely provide good information for conservation planning. This means that it is essential to identify sampled specimens to species level. Where this is not possible, morphospecies can be used, bearing in mind that they have far less currency than actual, identified species. In biodiverse localities where only some species are identifiable, it remains useful to use a mixture of clearly recognizable morphospecies alongside identified species for purposes of conservation planning.

2 Taxonomy and curation of insects

2.1 Introduction: essential planning Many insect species are unknown to science. Even species that have been described are often difficult to recognize and identify. This is the basis of the taxonomic challenge raised in the previous chapter. This dearth of knowledge is a major consideration for much of what we do in insect ecology and conservation. Whereas surveys of birds or mammals can often be undertaken without collecting and keeping specimens, this is usually not the case for insects. Apart from some well-known groups, such as certain butterflies and dragonflies, generally it is necessary to capture and preserve voucher specimens for identification by a specialist. Ecological insect studies often involve generalized collecting techniques that automatically capture a whole suite of species, including many non-target taxa, which may not be an intended part of the study. Together, these non-target taxa are known as bycatch. This generalized approach has associated ethical and practical problems, as discussed in the previous chapter, and presents risks to rare and threatened species. Indeed, the focal taxa of your study may well represent only a small proportion of the total insect sample, with the remainder never being used to advance knowledge. Even targeting only a single species may involve accumulation of bycatch. This remainder, rather than being discarded, should be kept for later study, perhaps by someone else in the future. This has particular value when, for example, a remote location has been surveyed or unusual specimens have been captured. Collecting and sorting samples is very time-consuming and laborious, and it is essential that collation of samples is planned with much thought and care, both for ethical reasons and for efficient use of time. In other words, you need to develop a strict sampling protocol. When sampling is done strategically and efficiently, important ecological information on spatial distribution and temporal

18 | Taxonomy and curation of insects

Collection parsimony

Nontarget taxa (bycatch)

Total sample

Focal taxa or individuals; contingency collecting

Fig. 2.1 Most collection techniques capture a range of taxa (the total sample) beyond those being targeted (the focal taxa, or other individuals e.g. functional types, size classes). These non-target taxa constitute bycatch that should be kept for possible future study. Sometimes it is appropriate to engage in contingency collecting (see Chapter 1), where taxa and individuals (which may or may not be the targeted, focal taxa) are collected for future studies, e.g. morphometrics or DNA. For both ethical and ergonomic reasons, insect ecologists and conservationists should always practise collection parsimony (see Chapter 1), where only those individuals essential to fulfil the project are collected.

activity patterns will be obtained. At the outset, it is important to establish (1) which insect group(s) is/are to be studied, and (2) the level of taxonomic interpretation that is needed to answer your research question. The position and numbers of samples and the sampling extent (see Chapter 3.3.2) to be examined must be clearly defined. As discussed in Chapter 3, the aims of the sampling programme (i.e. what you want to show) are integral to this. In short, your sampling should as targeted as possible. This theme is a major component of Chapter 3, but is introduced here to emphasize the importance of essential planning for interpreting insect samples, that, in turn, must be properly referenced in a collection (the subject of this chapter). In contrast to generalized trapping of many species at a time, conservation surveys may be focused on particular species. These are often conspicuous or distinctive taxa, which need not necessarily be killed for identification. Indeed, for single-species conservation studies, it is often essential not to kill or even damage them—for example, by handling during mark–release–recapture studies in population dynamics (Chapter 7). Murphy (1989) raises the sobering question ‘Are we studying our rarest butterflies to death?’, which summarizes the dilemma that we need data on threatened species but we must gather them without harming the individuals or the population.

Selection of insect groups for study | 19

In this chapter, we outline some ways in which the insects in samples can be assessed, and explore the importance of knowing the insects you are studying. As well as being able to recognize each insect species (or equivalent) reliably, knowledge of its ecology and seasonal pattern of development helps you to decide how and when to sample. We include discussion of the selection of which groups to concentrate on to gain good information (section 2.2); the extent of taxonomic interpretation needed (section 2.3); possible substitutes for taxonomic species (section 2.4), necessitated by lack of taxonomic knowledge in diverse faunas; approaches to rapid assessment using less specialized labour (section 2.5); and the importance and care of collections of voucher specimens as reference points for your study section 2.6). It may seem strange that this chapter has come before Chapters 3 and 4, which are on sampling procedures and methods of catching insects. We do this to emphasize that the aims, scope, and outcomes of your project must be carefully thought through before you start collecting.

2.2 Selection of insect groups for study Inventories that are aimed at sampling all taxa in a particular area (all-taxon biological inventories, ATBIs), although noble in intent, are usually impractical. A more practical approach is to survey particular taxa (focal groups). Simply selecting groups because they reflect the individual interests of participating scientists requires caution, because ‘favourite’ taxa may not provide the best possible information for the particular ecological or conservation research question you are asking. Focal groups ideally should be easily sampled, reasonably diverse, have acknowledged or suspected indicator or other surrogacy values, and be sufficiently well understood taxonomically that many or most of the species can be recognized and identified. Thus, water quality is commonly appraised on an EPT index. This involves a focus on the larvae of Ephemeroptera (mayflies), Plecoptera (stoneflies), and Trichoptera (caddisflies) (Resh and Jackson 1993), many of which can be recognized to species level through availability of user-friendly keys and illustrated handbooks. All of these taxa may change in abundance as a reflection of changing water quality. In geographical areas where these EPT taxa are less well known, adult Odonata can be excellent surrogates (Smith et al. 2007). It is entirely natural to gravitate toward the relatively few ‘well-known’ groups of insects for assessments and ecological analyses, both for ease of taxonomic interpretation of material collected and for the amount of information they consequently furnish. Indeed, many terrestrial insect surveys specifically select

20 | Taxonomy and curation of insects

certain focal groups such as ants or carabid beetles. Groups such as these are chosen because (on the basis of substantial literature evidence) their species richness and assemblage composition may be linked in some predictable way with habitat quality, as well as to changes in that quality in response to anthropogenic disturbance. Thus, the richness and ecological diversity of ants in Australia has led to delineation of a series of functional groups, whereby ants can be allocated to a particular ecological category, and the balance between the groups changes as conditions change in the wider environment (Andersen 1995b, Majer et al. 2004) (Boxes 2.1 and 2.2). The groups are founded on genera or other higher taxa (such as major species complexes), so obviating the need for species-level identifications (and allowing for inclusion of undescribed species), and the scheme has led to ants becoming the most frequently used insect ‘indicator taxon’ for terrestrial ecosystems in Australia. In many parts of the northern hemisphere, carabid beetles have been shown to be a useful focal taxon (Niemelä et al. 2000), but in many ecosystems in southern Africa and Australia (but see Horne 2007) their diversity is comparatively low and their taxonomy poorly understood. There is thus geographic variation in the suitability of particular taxa, and it is important to ensure that the chosen focal group is appropriate for the geographical area you are working in (see Chapter 10).

Box 2.1 Groups of ants used in environmental change assessments Functional groups of ants (Hymenoptera: Formicidae) are listed below, with comments based on Majer et al. (2004). Their representation in Australia, North America, and South Africa is shown in Table 2.1. 1 Dominant Dolichoderinae (DD). These behaviourally dominant ants are predominant in productive ant communities. The role is dominated by members of the subfamily Dolichoderinae. 2 Generalized Myrmicinae (GM). These ants are often abundant and assessed as subdominant to DD. They tend to have lower activity, smaller colonies, and smaller foraging territories, and are broadly distributed in relation to stress and disturbance. They tend to predominate in moderately (rather than high) productivity environments for ants. 3 Opportunists (OPP) are unspecialized ants, often with wide habitat distribution, and are poor competitors. They thus dominate on sites with low diversity, such as those where stress or disturbance limits diversity of other ants. 4 Subordinate Camponotini (SC) are based on the vast genus Camponotus, which tend to be behaviourally subordinate to DD.

Taxonomic knowledge | 21 5 Climate specialists (CS) comprise taxa with distributions centred on the arid zone (Hot Climate Specialists, HCS), cool temperate regions (Cold Climate Specialists, CCS), or the mesic tropics (Tropical Climate Specialists, TCS). 6 Cryptic species (CRS) are those tiny ants (mostly Myrmicinae or Ponerinae) that nest mainly in soil, litter, or decaying wood, and are essentially removed from ant competitive hierarchies. They tend to be most abundant in forests, and may be a major component of litter ant communities. 7 Specialist predators (SP) are medium-sized to large and strongly predatory ants. They tend to forage singly (the group was at one time called ‘solitary predators’) and have little competitive interaction with other ants.

The importance of knowing your insects extends well beyond simply being able to identify them. It also includes understanding their responses to habitat change and other environmental changes (such as climate change). In this broader context, insects become ‘tools’ in conservation assessment where they are used to indicate a change in the environment (see Chapter 10). Whether insects are the focal taxa of the conservation exercise, or whether they are just ‘tools’, accurate identifications and consistent recognition of the particular taxa involved is still necessary. In studies of single species, a ‘name’ helps to retrieve existing information on the taxon and, conversely a ‘wrong name’ yields misinformation (Thompson 1997). Studies focusing more on assemblages or communities, or even on the whole fauna of a particular site, require consistent recognition of many species to document and track changes in species richness and composition over time.

2.3 Taxonomic knowledge Even closely related species of insects, such as those in the same genus, may differ greatly in their abundance, basic biology, and responses to environmental change. Consistent recognition of species (or some equivalent working unit, as noted later) is the foundation for interpreting how a community works. However, many species cannot be named or recognized even by specialists, particularly in the tropics. Even for the best-known regional insect faunas, such as that of the UK, substantial proportions of very diverse groups such as parasitic Hymenoptera, Diptera, and many small beetles can be named only with much difficulty and with advice from taxonomic specialists (Barnard 2000). A field ecologist cannot assume that all insect groups collected in a survey can be identified reliably to species level, even in the better-documented places. Some form of triage (the selection of particular insect groups for study, disregarding others), however ethically

22 | Taxonomy and curation of insects Table 2.1 A key to groups of ant functional groups between Australia, North America and southern Africa. Major genera are listed in general order of ecological importance. Tropical rainforests are excluded because this habitat is present only in Australia. From Majer et al. (2004). See Box 2.1 for explanations of ‘groups’ Functional group Australia

North America

Southern Africa

DD

Dorymyrmex (bicolor gp) Forellus Liometopum Pheidole

Absent (except invasive aliens)

GM

OPP

Iridomyrmex Anonychomyrma Papyrius Monomorium (several) Pheidole Crematogaster Rhytidoponera Tetramorium Paratrechina Aphaenogaster

SC

HCS

CCS

TCS

CRS

SP

Camponotus Polyrhachis Calomyrmex Opisthopsis Melophorus Monomorium (several) Meranoplus

Notoncus Prolasius Stigmacros Monomorium (leae, kiliani gps) Podomyrma Oecophylla Podomyrma (some) Tetraponera Monomorium (some) Solenopsis (Diplorhoptrum) Hypoponera Myrmecia Cerapachys Pachycondyla Leptogenys

Monomorium (most)

Crematogaster Monomorium Aphaenogaster Myrmica Dorymyrmex (insana gp) Aphaenogaster Paratrechina Camponotus

Camponotus

Pogonomyrmex Myrmecocystus

Ocymyrmex Tetramorium (solidus gp)

Messor

Monomorium (setulosum gp)

Solenopsis

Messor Anoplolepis (some) Anoplolepis (some)

Formica (several) Lasius Leptothorax Prenolepis Stenamma Neivamyrmex Attini Pseudomyrmex Solenopsis (Diplorhoptrum) Acanthomyops Hypoponera Polyergus

Pheidole Crematogaster Tetramorium Lepisiota Paratrechina

Dorylus Myrmicaria Cataulacus Meranoplus Solenopsis (Diplorhoptrum) Plagiolepis Pachycondyla Leptogenys Plectroctena

Taxonomic knowledge | 23

challenging the concept may be, is almost inevitable in collating the material accumulated from insect surveys (New 1998). Taxonomic tractability is a very important practical aspect when selecting particular focal groups. Insects can be divided into three broad groups relative to taxonomic knowledge on them (New 1999): • Well-known groups. In these groups many of the species are named or recognizable, with illustrated keys or handbooks available to help identification. Unfortunately, few insect groups come into this category, although they include butterflies, dragonflies, and some groups of beetles (such as carabids and tiger beetles). A bonus of working with well-known groups is that improved taxonomic knowledge is usually correlated with improved biological knowledge. • Catch-up groups. These have a strong established taxonomic framework, perhaps to generic level, and many of the species have scientific names. Some identification guides may be available, but with significant gaps in coverage, and so they are not suitable for use by most non-specialists, because errors can easily be made. Such groups could usefully be made the focus for taxonomic studies because they could be converted to ‘well-known’ with relatively little work, and could then augment the portfolio of insect groups that could be interpreted meaningfully in surveys. • Black hole groups. These are the groups of diverse, small, and inconspicuous insects, for which taxonomic knowledge is relatively poor, and in which many species have not been recognized, diagnosed, or named. The taxonomic literature for these groups is mostly specialized rather than widely available as identification handbooks to non-specialists. The incompleteness of knowledge renders almost all identifications by nonspecialists unreliable. Nevertheless, many such groups have massive ecological importance. Disney (1986), for example, considered the values of flies and parasitic Hymenoptera in ecological interpretation to be high, but the groups are simply intractable at present, and can be employed only in very approximate terms. Likewise, poorly understood parasites, even if not numerically abundant, can have disproportionately important roles in ecosystem functioning (Hudson et al. 2006). The goals of your study must be very clear, so that you can select your focal group bearing in mind the extent of taxonomic expertise or support available. Indeed, pragmatic criteria will be fundamental to selecting and narrowing down the groups to be studied. For a freshwater insect survey in Australia, Cranston and Hillman (1992) selected taxa on (1) availability of specialist taxonomic

24 | Taxonomy and curation of insects

expertise to validate identifications, (2) availability of a good local identified reference collection, including earlier voucher material from the study site, and (3) acknowledgment by others that the groups have special merits as biological indicators. It is useful, wherever possible, to seek advice from more experienced workers or taxonomists, particularly where there might be doubt about the level of tractability of any group proposed for survey, and you should do this well before a study starts. A relevant specialist may, for example, be able to give advice on selection and adequate sampling of the insects and their subsequent treatment. Such a specialist is also likely to have substantial background biological knowledge and ability to recognize any especially interesting species in the material that you should look out for at your study site. There is also an economic consideration. If assistance is required for sorting, mounting, and identifying the insects collected, this should be costed realistically in planning the project budget. Also, there should be clear agreement with such collaborators on the intellectual property issues involved (such as coauthorship of papers), where material will be stored, and where any type material will be permanently deposited. It is not good practice to take specialist aid for granted, or to expect an expert to identify large arrays of inadequately processed or poorly preserved material without prior arrangement or subsequent acknowledgement. In many cases, taxonomic institutions are obliged to charge fees for identifications to cover the considerable time required. Some taxonomists however, may be very interested in your collection, and may request that they keep your material in exchange for identifications. This is usually a fair deal, but before embarking on it, you must clear it with your own institution to ensure that this is an acceptable arrangement. We discuss below (section 2.5) some of the ways in which samples can be interpreted without having to undertake formal species identifications. Nevertheless, it is always preferable to have material properly identified, as it then has much greater value for communication and for comparison with already published data (Thompson 1997). The history of applied entomology, in particular, includes many cases in which misidentification of key pests, or confusion between related or similar species, has hampered management programmes. Similar situations are also beginning to arise in conservation in species-rich areas of the world. For many insect groups, only the adult stage can be recognized and diagnosed with confidence. Most surveys of terrestrial insects are limited to adult insects. Yet immature insects often predominate in samples, but most of these have not been associated with corresponding adults by direct rearing, so cannot be identified reliably. Freshwater insects have a rather different bias, as the

Taxonomic resolution | 25

larvae of orders such as mayflies, stoneflies, and caddisflies in some geographical areas are well known and distinctive at the generic or species level, and occasionally may be more easily diagnosed than the adults. Nevertheless, very few surveys have compared adult and larval samples of insects in a given site or habitat, although Hawking and New (1999) compared the distribution of larval and adult dragonflies along an Australian river. Andersen et al. (2002) make the interesting observation that sampling just large-sized ants (those more than 4 mm long) reduced sampling times normally devoted to all ants down to only 10% and yet produced sound results. It may be possible to use subsets of taxa where appropriate and still get good results (that might not necessarily be simply large-sized individuals of other taxa). Indeed, this idea of subsets as surrogates is an exciting field of research in its own right and can enable substantial time saving when working at the critical and sensitive level of species, rather than at insensitive, higher-taxonomic levels, such as families (see next section). However, the conclusions drawn from using species subsets has been little tested (McGeoch 2007). Caution is needed so as not to make unjustified assumptions.

2.4 Taxonomic resolution One aspect that can help overcome lack of species level information is to identify material to a lower resolution, i.e. to only higher taxonomic levels, such as by order, family, or genus, as well by functional groups, as mentioned for ants above. It is almost inevitable that such steps reduce the detail and value of the biological information obtained. Studies of insects categorized to only ordinal level are often difficult to use because such umbrella-groupings are virtually ubiquitous, and little information is conveyed. Classifying insects to orders is similar to classifying vertebrates simply as ‘birds’, ‘mammals’, ‘reptiles’, and so on, with each category containing a multitude of ecologically disparate forms. Such approaches are deceptively attractive, because they are simple and cheap to pursue, require little specialist discrimination, and little training to provide consistent results. A broad picture may indeed be gained, but the finer brush-strokes needed for refined management and critical assessment of diversity are almost inevitably missing. Danks (1996) notes that even family-level separations in insect samples are generally inappropriate because of extensive speciation at lower taxonomic category levels, which is often where there is the most subtle ecological sensitivity and discrimination. Any such approach using higher taxonomic groups, or focal groups, as surrogates for wider biodiversity is part of the general trend to seek rapid

26 | Taxonomy and curation of insects

biodiversity assessment (RBA) methods to gain information at a lower cost than when undertaking more complete assessments. Ward and Larivière (2004) discuss four main categories of RBA involving insects, as follows: • Sampling surrogacy (reduced sampling intensity), such as trapping for shorter periods or reducing the number of individual trap units. This can be very attractive in reducing later costs, but care is needed to ensure that the regime remains scientifically rigorous (Chapter 3). • Species surrogacy: as noted above, this is where higher groups are substituted for species-level analyses. • Taxon surrogacy: the use of morphospecies (below) rather than formally identifying all components of the samples. • Taxon focusing: the selection of particular taxonomic groups as surrogates of wider biodiversity. The practical savings in time and costs gained from such broad approaches may still allow worthwhile assessment of complex insect samples, and set a foundation for more detailed, perhaps later, taxonomic analysis. Tests of the value of higher-level surrogacy have provided mixed results, so that it is usually not clear whether, for example, generic diversity strictly parallels species diversity within an insect group across samples or sites (see biodiversity indicators, Chapter 10). One instructive example involves ants in Australia. Most of the hundred or so genera (Shattuck 1999) can be recognized reasonably easily from user-friendly published keys, but many species identifications are precluded by lack of recent revisions and the presence of many undescribed taxa. Generic and species richness are sometimes closely correlated (Andersen 1995a), although this may sometimes result from the presence of many species-poor genera in samples (Neville and New 1999). Because ant functional groups (Box 2.1 above) are specified at levels above species, the ecological information obtained from these functional groups is largely valid. Also for the objectives involved high taxonomic resolution (identification to species level) is not necessary. Any such surrogacy measurement should be employed only on the basis of clear understanding and initial testing of the exact relationship between the higher and lower taxa (New 1999). Without this, serious errors may occur. For example, when there is an environmental disturbance rare and threatened species are often lost, and common and widespread species are gained or retained. In this case, if assessments are done at higher taxonomic levels this effect would not be detected and the approach would be counterproductive for conservation assessment. In summary, we strongly recommend that you do your study at the species level, rather than at higher taxonomic levels.

Morphospecies and parataxonomists | 27

2.5 Morphospecies and parataxonomists Most of the costs associated with insect sampling are incurred by the laborious sorting and identifying. Yet these must be done carefully to ensure consistency, which is important when quantifying samples, or comparing sites, or comparing the same sites over time. With a shortage of taxonomists relative to the diversity of insect life, it is imperative to use alternative methods so as not to slow the urgent task of insect diversity conservation. To this end, two tactics are widely employed in insect studies: use of non-named but consistently recognizable entities (morphospecies; section 2.5.1, Fig. 1.5), and employment of biodiversity technicians (parataxonomists) to replace or precede more specimen identification by specialists (section 2.5.2). These two methods commonly go hand in hand.

2.5.1 The morphospecies approach The term morphospecies (see Figure 1.5) is used to denote a consistently recognizable entity thought to be equivalent to a species, but not named formally as such. The underlying principle of their use is that morphospecies are (1) deemed to be true surrogates for ‘real’ species, and so (2) may be used to classify and represent insect diversity without compromising accuracy, reflecting that the most commonly used measure of diversity is species richness (the number of species). Morphospecies can be separated on any character or suite of characters that are sufficiently constant to distinguish amongst the entities involved. A variety of equivalent terms, such as recognizable taxonomic unit (RTU) or operational taxonomic unit (OTU), have been used, and are essentially synonyms (however, as a precaution, you should read the background definition to any such term you come across in a particular research report). Use of any such entity depends on consistency of recognition, and that each morphospecies contains all the representatives of a given type, and only those individuals. Sorting is therefore subject to the same constraints as for named species with neither splitting (the excessive: there should be division of entities on minor or non-significant variation to inflate the number of species or morphospecies present) nor lumping (the amalgamation of different forms through failure to recognize consistent differences, so lowering the number of real taxa). However, it is always better to err on the side of splitting, because, should you find at the end of your sampling that what you thought was two species (or more) now turns out to be one, it is easy to combine figures in your data matrix. But if you have one species and it turns out to be two or more, it means that you will have to sort through all your samples again to separate the individuals into their respective species.

28 | Taxonomy and curation of insects

It is important to recognize the advantages and limitations of morphospecies. Different groups of insects pose different problems for consistent recognition. In one comparative trial using ground-dwelling forest invertebrates, Oliver and Beattie (1996) found that ants and some beetles showed very high correspondence between morphospecies and species. In contrast, other families of beetles were much more difficult to identify, and morphospecies:species equivalence was much lower. The morphospecies approach allows inclusion of poorly known taxonomic groups into assessments of diversity. The ideal, of course, is that a morphospecies is fully equivalent to a real taxonomic species in such estimations. You may well need guidance and training to ensure that this ideal is attained. This may involve some preliminary sorting to gain practice before embarking on a full sorting exercise. Constant referral to voucher specimens (i.e. reference specimens as representatives of every different entity that you distinguish) is crucial at this stage. When working in biodiversity-rich and poorly studied areas, it is highly unlikely that all the specimens collected will be able to be named, even by specialists, and so it makes very good sense to build a wellcurated collection of voucher specimens, which may contain both identified and unidentified (morphospecies) specimens. Clearly however, one must also recognize the long-term limitations of using morphospecies: for example, it precludes comments on endemism, threat levels to specific species (e.g. Red Listing), and detailed autecological studies. Inevitably, separation of species (or morphospecies) based on morphological characteristics reflects a typological approach to species distinctions, and may mask unknown levels of variation. As Heywood (1994) emphasized, a ‘species’ may consist of many populations with differing genetic constitutions and adaptive potentials, and the lumping vs splitting dilemma becomes important in assuring reasonable consistency in sorting. There may indeed be a temptation to produce long species or morphospecies lists as inventories, because of the political message of high diversity they convey, but in other situations the emphasis may be on ‘how few species’ rather than ‘how many species’ are present. In most cases, quality control over the consistency of recognition can be improved in two main ways: • Continued cross-referencing to a complete set of the voucher series, sorted and accumulated progressively as the work proceeds. • Progressively accumulated notes, sketches, and digital images for each putative morphospecies and its diagnostic features (which would not be a formal taxonomic description), providing a working tool for consistent recognition.

Morphospecies and parataxonomists | 29

Such accounts would also be supplemented by the voucher collection. These records are important, because there are differences in perceptions among different researchers. Good quality multidimensional digital images can help overcome these observer differences. The use of morphospecies in your study should be considered very carefully. Some of the problems and possible limitations of their use are summarized by Krell (2004), who emphasizes the need for effective quality control. However, that summary is naive in terms of practical conservation sampling, especially in the tropics. It overlooks the real value of morphospecies as stepping stones to later taxonomic verification. It also ignores the fact that in many parts of the world insect species simply cannot be named because there are so many species, many of which may well be undescribed, or there is no scientific specialist available to do the job. A final point is that where the study covers a range of well-known, catch-up, and black hole groups, you may, in your data matrix, be able to insert taxonomic names for the well-knowns and morphospecies names for the catch-ups and black holes (e.g. Buprestidae sp. 1, Buprestidae sp. 2, …). This is a particularly useful approach where you are working in lesser-known geographical areas such as the tropics.

2.5.2 Parataxonomists The term parataxonomist was used initially for technical staff employed in Costa Rica under that country’s developing national biodiversity inventory (InBIO) to sort samples of insects and other organisms to morphospecies. Parataxonomists are usually employed on very large projects involving teams of researchers. The approach helps overcome lack of specialist capability or availability (Janzen 1997) by employing and training local people to collect, process, and undertake preliminary identifications of insects based on morphological recognition. This approach does not solve all the problems of sampling, because, as pointed out by Basset et al. (2000), deployment of a team of parataxonomists can produce a huge amount of data but it still needs to be adequately analysed and written up by appropriately skilled researchers. Also, where parataxonomists are used, there must be standardization between individual workers, otherwise the value of parataxonomy breaks down. Problems in sorting may arise from factors such as sexual dimorphism (so that males and females of the same species are treated as different taxa); polymorphisms (various ‘forms’ of the same species); small size (obscuring small morphological differences); presence of very similar species; separation or amalgamation of different instars or growth stages; as well as individual observer traits (such as colour blindness). Several studies have indicated,

30 | Taxonomy and curation of insects

for instance, that an individual’s sorting acuity can lessen over a period of several hours of work as they become bored or careless with increasing tiredness. Nevertheless, with adequate initial training and continuing supervision and monitoring, highly reliable results can be obtained. For very diverse samples of tropical insects, in particular, Basset et al. (2000) noted the benefits from their activities as including (1) increased efficiency of year-round sampling, (2) rapid processing of high quality specimens at low cost, (3) enhanced interpretation of local ecological information associated with the specimens collected, and (4) increased local interest in biodiversity. In short, subject to good quality control, their activities can markedly enhance appraisal and understanding of insect surveys. Essentially, employment of non-specialists has accelerated the collection and processing of specimens to lead toward inventories of rich insect faunas in countries with little resident entomologist expertise. Even in entomologist-rich parts of the world, non-specialists have vital roles to play (Goldstein 2004). These approaches are often advocated as contributions to rapid biodiversity assessment (RBA), sometimes without adequate emphasis on the need for consistency. The practical balance needed is that between (1) reducing costs and time through use of non-specialists and imprecise species categories, and (2) incurring increased error risk and frequency through reduced specialist involvement. Parataxonomists are not an alternative to professional taxonomists, but enhance their capacity and capability (Lewis and Basset 2007). Wilkie et al. (1999) proposed a basic sorting protocol, summarized in Fig. 2.2, in which feedback on quality control is provided as training at several stages: during sorting invertebrates to class, to order, and to morphospecies levels. This form of process control sampling was regarded as the better general strategy to adopt, not least because it is proactive rather than reactive, and serves to minimize errors during the entire sorting process.

2.5.3 Voucher collections Such controls need, and demonstrate the practical values of, progressive accumulation of a voucher collection, i.e. a reference set of the morphospecies distinguished during sorting. A voucher collection (or reference collection) is built so that the individual designated as ‘morphospecies no. 16’ (for example) has a reference point against which other individuals can be compared. In many ways, a voucher series serves the same purpose for an ecologist as do type specimens for a taxonomist. Both are the ultimate reference points for accuracy, and vouchers must therefore be preserved, documented, and archived safely (e.g. see below) for future study. The uses of voucher collections include: (1) checking

Bioinformatics | 31

identifications as systematic knowledge advances, (2) detecting changes when re-sampling sites at some later date, (3) validating any reinterpretation or reanalysis of the data, and (4) comparing sites, such as those under different land use (Lee et al. 1982). Without such vouchers, errors of identification and interpretation may never be detected or verified. The responsible deposition of voucher collections, preferably in a state or regional museum where proper curation (section 2.7) can be assured, should be an integral part of planning an insect survey, and the costs of this (and for subsequent maintenance) should be included in the project budget. At present, many investigators neglect this aspect of their surveys. The concept of virtual vouchers (see below) is an invaluable adjunct in sample processing, because of the easy availability of high-quality comparative images. However, it does not replace the need for actual voucher specimens that can, if necessary, be dissected or otherwise more closely examined in the future. A similar responsibility applies to the bulk samples from surveys. Danks et al. (1987) referred to these as ecological collections, currently of unknown value for future investigation and for monitoring ecological changes at the sites from which they were derived. These ecological collections should be clearly and permanently labelled and stored responsibly as long-term archival collections, for genetic studies as well as for traditional taxonomy. Scudder (1996) expressed a much more widespread view, that on both ethical and more pragmatic grounds such collections should not be discarded: not only have the insects been killed, but there may have been substantial investment in obtaining them. In some cases, samples may be from remote localities that may be very difficult to revisit. From the taxonomist’s point of view, such places may be of special interest in yielding unusual forms of variable taxa or members of sibling groups important for demonstrating patterns of evolution or endemism. Indeed, the deliberate accumulation of ecological collections may be a wise investment in documenting our invertebrate biodiversity. As Yen (1993) put it, ‘if this kind of collection had been made more widely in the past, we would probably have many more answers to conservation management issues—but it is not too late to make ecological collections for use over the next century’.

2.6 Bioinformatics Biological informatics technology, or bioinformatics, is the application of information technology to solving biological problems, and it is becoming an important theme in the description and identification of organismal biodiversity. Many aspects of technology are rapidly becoming tools or components of

32 | Taxonomy and curation of insects Field collection Curation

Training

Feedback on errors

Sort invertebrates to class

Data entry on form

Quality control

Preparation of identification guides

Curation

Training

Feedback on errors

Sort focal taxa (insects and arachnids) to order

Data entry on form

Quality control

Preparation of identification guides Curation

Training

Sort focal taxa to 1 morphospecies per vial

Feedback on errors

Data entry on form

Quality control

Preparation of identification guides

Arrange vials by morphospecies per vial

Fig. 2.2 A basic laboratory sorting protocol for insect specimens. Intermediate levels (e.g., sorting to family) may be inserted in the appropriate location using parataxonomists. NB: Each new level inserted has to have its own step of quality control and training (from Wilkie et al. 1999).

insect sample determination, in providing additional viewpoints for assessing diversity, and streamlining transmission and management of the information accumulated (Johnson 2007). The following section outlines ways in which bioinformatics is applied.

Bioinformatics | 33

2.6.1 Virtual voucher collections In almost every case, the voucher specimens are dead insects preserved for study by the methods outlined later in this chapter. However, alternative (or additional) approaches are gaining popularity. The development of accessible, high-quality digital photography, and accompanying software that allows photographs to be downloaded and transmitted within minutes, has opened up the potential for photographic or virtual vouchers as photographs in which recognition or diagnostic features are displayed clearly. Oliver et al. (2000) showed how such systems may be used in assessments in ways that can be of practical value to conservation planners and land managers. Local virtual voucher collections (as digital image compilations) of species or morphospecies may be established, including images of the morphological characters by which each morphospecies is recognized and diagnosed. Such collections also facilitate rapid transfer of information, so that different laboratories and workers can use the same baseline for assessments, and so achieve fully standardized and comparable results. Importantly, such systems should use accessible and generally affordable software to improve the speed and accuracy of data acquisition and manipulation. Barcode technology is used for sample and specimen tracking, and a high-resolution image-capture system is used for reference specimens and specific characters as each new entity is added. This virtual voucher collection, with associated taxonomic information, is used for onscreen recognition/identification of species, with potential to achieve wide usage on the World Wide Web as a directory of biodiversity (such as the Web of Life, http://tolweb.org/tree). Macadam (2006) notes the technical approaches to achieving high-quality voucher photographs. Importantly, however, he emphasizes that a voucher photograph can never be a full substitute for the preserved insect, but is a complement to more traditional vouchering. In his example and words ‘there is no reason why a voucher photograph of a live moth should not accompany a set specimen in a collection’. The software package Biota: The Biodiversity Database Manager is designed specifically for biodiversity databasing and includes a facility for image storage.

2.6.2 Sample tracking Allocation of a unique barcode label (see section 2.6.3) to each sample and subsequently sorted or mounted specimen increases the speed of data acquisition and helps raise quality by eliminating errors due to label transcription. Onscreen identification (1) is fast, (2) can be undertaken simultaneously by more than one person, and (3) obviates the need for repeated handling of real,

34 | Taxonomy and curation of insects

fragile reference specimens. It is also claimed to increase accuracy and consistency of specimen allocations, and transmission of images may help in gaining advice from specialists far from the survey laboratory. A taxonomist could in theory, and increasingly in practice, work more from images accumulated from collections throughout the world, with digital images of holotypes and other critical museum material reducing the need for posting (and risking) such valuable specimens. The usefulness of this approach depends on the quality of the images and auxiliary information, as well as the costs of obtaining them.

2.6.3 Taxonomy using barcoding Molecular data, particularly through the use of DNA barcoding, is a powerful tool for detecting so-called cryptic species (insects that resemble each other very closely so that they cannot be separated on morphological grounds alone). Molecular data thus helps to estimate species richness, to indicate intraspecific variation, or to delimit entities as biologically and taxonomically distinct (Wilson 2008). The process involves the sequencing of a single gene region, generally a 688 bp piece of mitochondrial cytochrome oxidase subunit-I (COI or coxI, for short). Cheap, rapid processing is available to do this. Specimens are not destroyed in this process, and analyses can be made from a single leg, or small part of a wing, so that the remainder of the insect can be kept in a normal collection, with suitable cross-labelling to the genetic material. However, specimens killed by chemicals such as ethyl acetate may have degraded DNA. To prevent this they should rather be killed by freezing or immersion in absolute alcohol. The barcoding approach has already proved its value in separation of sibling species of pest insects. Perhaps the largest and most pertinent example to note here involves an inventory of the larger Lepidoptera of a major reserve in Costa Rica, a region where many of the species have not been described, and based on a survey that has continued for almost 30 years. It also illustrates the critical roles played by parataxonomists (section 2.5.2) in such exercises. Since 1978, every caterpillar found in the forest of the Guanacaste Conservation Area has been kept (background in Miller et al. 2006), given a unique voucher code, photographed, and reared to produce an adult or its parasitoid(s). Adult Lepidoptera are killed by freezing, and genetic analysis is based on legs of selected specimens. By the end of 2004, the project had accumulated 264 370 individual records, representing more than 1000 species, and providing unique insight into the richness and variety of a tropical insect group (Janzen et al. 2005, Box 2.2). Most similar projects will be on a much smaller scale. For example, a putative single species of skipper butterfly was shown using DNA characteristics to comprise 10 distinct species (Hebert et al. 2004).

Specimen treatment | 35 Box 2.2 The process and background used to provide an inventory of Lepidoptera in Costa Rica based on DNA barcoding 1 Foundation: many individually vouchered and databased specimens have been accumulated as museum material over two decades from all main ecosystems of the Area de Conservacion Guanacaste. 2 Each insect voucher specimen is available automatically as a barcode voucher specimen. 3 The frozen and then oven-dried specimens have not been processed by methods known to destroy their DNA. 4 Each adult insect reared has three pairs of legs, and one member of a pair can be removed for sequencing, with ‘reserves’ available if needed. 5 The specimens are already identified to some level, often putative species, by more conventional taxonomic procedures, before entering the barcoding process. 6 When barcoding generates taxonomic questions, the inventory process can be modified to generate more specimens of the taxon in question, although this will of course involve a time lag maybe of up to a year through the process of discovery and rearing. 7 All species examined are either sympatric within the area or, if restricted to different ecosystems, parapatric at the meetings of those ecosystems over distances of a few hundred metres. 8 The specimens being studied are usually in excellent condition, as freshly reared rather then flight-worn adults. 9 Because they are all reared, it is often possible to know whether a pair of specimens is likely or actual siblings, and even to use barcodes of siblings and parents to examine intra-population variation and to confirm the accuracy of sequencing.

(Janzen et al. 2005)

2.7 Specimen treatment Processing insect samples can be much more expensive than collecting them in the field, because of the large numbers of specimens accumulated. One published estimate suggested a sampling:processing cost ratio reaching 1:40. Thus, capturing and killing insects implies both a duty of care for the material, and the ability to track every specimen from a sample to its original source by adequate documentation (such as labels) and to keep it in good, examinable condition. Where temporary labels with information such as ‘trap 46, transect 7’ or ‘grassland pitfall trap

36 | Taxonomy and curation of insects

no. 12’, have been used, it is essential to replace these temporary codes immediately with labels carrying information more meaningful to others. Similarly, using simple codes in notebooks can also carry risks (notebooks can get lost), and so, whether labels or notes, it is far better right from the start to use full information that is meaningful to any other researcher. This is one of the advantages of digital data loggers, where at least spatial information (such as geographical coordinates) is automatically recorded. In remote areas, where local place names are few and far between, and can change as the political climate changes, it is essential to record exact spatial coordinates. With the widespread availability of inexpensive geographic positioning systems (GPS), these data should progressively be collected as part of normal good practice. The data still need to be downloaded, and where appropriate, added to the actual specimens. In short, the aim is to produce a voucher collection that is informative and meaningful to any future generation of researchers. At the time of retrieving each sample from the field, it should be provided with fuller information, in addition to any working code, with such labels placed routinely into each container of specimens. There are practical issues when doing this. For example, where the specimens are in alcohol, it is important to place the information inside the bottle, not outside, from where it may eventually peel off. Labels should be written in pencil and not in ink because some types of ink dissolve. Computer-generated labels on thin card can be prepared in bulk in conjunction with planning the sampling programme, so that each sample has a specially prepared label awaiting its preservation. Ensure that the ink is not soluble in the alcohol or in other preservatives (some workers recommend baking the labels in an oven at 120 °C for half an hour before use). A type size of 4–6 points for individual vials, and larger labels in 10–14 point type for bulk containers are generally suitable. Each subsample must have the same locality and ecological data on labels as the parent sample. Very careful attention should be paid to the type of bottle or vial (soda glass, because plastic is not suitable for long-term storage) used for storage of wet specimens (i.e. those in preservative). The bottles must have good seals that must not perish when in contact with the preservative, and must prevent evaporation. Bear in mind also that when the specimens are sent to a specialist via airmail the preservative must be drained, otherwise the bottles will not be handled by the air freight company. After the preservative (usually 80% alcohol) is poured out, the specimen must be plugged in the bottle with preservative-moistened cellulose wadding (as used for wound dressings in hospitals), and not with cotton wool that tends to snag the legs and antennae. The packaging for dispatch must also be labelled for customs purpose with appropriate wording such as ‘Preserved scientific insect specimens, of no commercial value, and without free inflammable liquid’.

Specimen treatment | 37

Details of label data will differ slightly for different purposes, but should normally include: (1) country; major political entity (state, territory, region); (2) name of nearest major settlement (with distance and direction from this), or name of national park or reserve (if relevant); (3) latitude/longitude or other recognized national grid reference or GPS reading; (4) major habitat; (5) collecting method (with any sample number, if relevant); (6) date of collection (day/month/year, cited as ‘10 March 2007’or ‘10.III.2007’—not ‘10.3.2007’); (7) name of collector (Fig. 2.3). These label cards are usually about 15 × 25 mm. The country name is necessary because many specimens are exchanged between specialists or sent to specialists overseas for identification, with their eventual home in museums far from their point of capture. In the absence of a country name, a specimen simply labelled ‘Victoria’, for example, could be derived from a state of Australia, a region of western Canada, East Africa, or several other widely separated places. The sequence of date notation is recorded as 10.III.2007 (or even better, with the month in full), because this is unambiguous to all nationalities, which is not the case with 10.3.2007, which could be interpreted as either 10 March or 3 October. If many samples are to be processed, and to remove transcription errors, barcode labels can be used, as an innovation from RBA approaches (section 2.4). The ideal label should allow the reader to return to the collection site, and use the same sampling method at the same season to re-sample there.

Fig. 2.3 An enlarged example of a data label that has been prepared to go on the insect pin just beneath the specimen (see Figure 2.16). Such labels usually measure about 25 ⴛ 15 mm. If there is not enough space on a single label, two labels can be used with one placed directly below the other on the pin. The second label usually contains the ecological or habitat information.

38 | Taxonomy and curation of insects

2.7.1 Sequence of steps for treating insect samples The following sequence outlines the major steps needed to process insect samples, be they single species or many species. Single-species studies may impose additional demands, such as needing specimens for genetic analyses, requiring particular storage or transport modes, but the procedures are otherwise similar. Many specialist taxonomists have developed their own preferred techniques, such as slide-mounting and dissections, for close examination of the insects they study. This means that it is essential that you contact the specialists ahead of time to make sure that you supply them with the material prepared in the right way. More detailed information is readily available (Upton 1991; Uys and Urban 2006). Although most insect sampling involves handling and identification of dead insects, it is sometimes necessary to cage and transport living insects for laboratory study, or experiments, establishment of breeding stocks, translocations (Chapter 8) or other purposes, such as rearing larvae to adults to facilitate recognition. Living insects may suffer or die from many causes, for example if: (1) they are exposed to temperature extremes, (2) deprived of suitable food, (3) overcrowded in small containers, or (4) not sufficiently confined so they can damage themselves on cage sides. More rarely, long-term travel, such as by air, may be needed, and it is then wise to arrange any documentation needed well in advance to avoid possible delays in quarantine or customs. The essential steps in preparing insect specimens for examination are described below.

2.7.1.1 Preservation Techniques for preserving and maintenance of insect specimens have been developed for well over a century and are described in a wealth of texts (see Oldroyd 1968, Smithers 1982, Upton 1991, Uys and Urban 2006, Cooter and Barclay 2006). Table 2.2 summarizes many of these standard and well-tried methods for each insect order, and the following notes augment this. First, the specimens must be killed humanely, then preserved in a suitable way, prepared for study as necessary, and finally labelled and stored responsibly. Studies for conservation or environmental assessment often involve taking of bulk samples by various types of traps, while surveys for particular species or guilds (such as pollinators) may involve collection of just a few individual insects. Bulk insect samples, normally of recently captured or killed insects, are commonly collected passively and directly into liquid preservative (as in pitfall traps, Malaise traps, or suction traps; see Chapter 4). Actively caught insects (with

Specimen treatment | 39 Table 2.2 Recommended preservation methods for different orders of insect. Formulations of fixative chemicals as in Table 2.3 Order

Methods

Archaeognatha

All stages: fix and preserve in 80% ethanol

Thysanura Ephemeroptera

As above Fix all stages in Kahle’s, Carnoys, or FAA and preserve in 80% ethanol: some workers recommend adding 5–10% glycerol. Adults can be pinned through centre of thorax Odonata Larvae: kill and preserve in 80% alcohol; exuviae in alcohol or glued onto cards. Adults killed and colour-fixed in acetone, and once dry, placed in transparent envelopes Blattodea Preserve dry: pin through centre of metanotum; usually do not set, but left wings can be spread if needed Isoptera Collect and preserve in 80% ethanol; series of all castes should be collected wherever possible to facilitate identification Mantodea Pin between base of wings. Extend one foreleg so spines on femur and tibia visible; spread left wings of vouchers. May be necessary to ‘gut’ large specimens, by slitting along the abdominal pleural membrane and removing contents with forceps (avoiding damage to sclerotized structures); the abdominal cavity can then be wiped dry, dusted lightly with talcum powder and boracic acid powder (3: 1), and a small wedge of cellulose wadding or tissue inserted before specimen is pinned and set Mantophasmatodea As above, pin through centre of thorax Phasmatodea Pin through base of mesothorax; gut large specimens as above. May be useful to fold legs and align antennae along the body, to save storage space. Spread left wings of vouchers Orthoptera Adults: pin vertically through prothorax, to right of midline; spread left wings of vouchers. Soft-bodied species and nymphs: fix in Pampel’s and preserve in 75–80% ethanol Dermaptera Adults; pin through right elytron or equivalent position to right of midline of mesothorax. All stages: Pampel’s and alcohol as for Orthoptera Embioptera Winged adults may be pinned through centre of thorax and wings set. Otherwise: all forms in 75–80% ethanol. Grylloblattodea All: 80% ethanol Zoraptera As above Plecoptera Adults: pin through centre of thorax and set. Otherwise, all stages: fix in FAA and store in 80% ethanol. Hemiptera (Heteroptera) Large adults; pin through scutellum to right of midline; smaller specimens: stage or card point dry. Spread wings of vouchers when necessary. Nymphs: 80% ethanol (Auchenorrhyncha)

Large adults; pin through scutellum to right of midline; smaller specimens: stage or card point dry. Spread wings of vouchers when necessary. Nymphs: 80% ethanol

40 | Taxonomy and curation of insects Table 2.2 (continued) Order

Methods

Coccoidea

Males: collect and preserve in 75–80% ethanol. Females: collect and preserve in lactic–alcohol, to facilitate treatment for later slide-mounting. Scales and mealybugs; dry and card as necessary

Aphidoidea

Collect into 80% ethanol or lactic–alcohol; may be useful to have some specimens in a cytological fixative

Aleyrodoidea

80–95% ethanol

Psylloidea

Adults: mount on card points or preserve in 80% ethanol; may be necessary to dry-preserve galls and lerps on foliage

Phthiraptera

80% ethanol, probably for later slide-mounting

Psocoptera

As above

Thysanoptera

Fix in 60–70% ethanol, probably for later slide-mounting

Megaloptera

Adults: pin through centre of thorax and spread wings, at least on left side of vouchers. Can fix all stages in FAA or 80% ethanol for storage in the latter

Neuroptera

Adults: pin representatives to right of midline through mesonotum, spread wings of left side or both sides. Others, and early stages, fix in FAA or Carnoy’s and store in 80% ethanol

Raphidioptera

As above

Coleoptera

Large adults: pin through right elytron about one third from anterior and one third from midline; usually not set. Smaller adults: stage, or mount on card rectangles or points. all stages in 85 % ethanol, after KAA or Carnoy’s

Strepsiptera

Males can be carded, but more usual to preserve all stages in 80% ethanol; if host available preserve this for association

Mecoptera

Adults: pin through metathorax to right of midline; wings may be spread. Others and early stages in 80%% ethanol after KAA

Diptera

Larger adults: pin to right of midline of mesothorax. Small adults, stage. Larvae: fix in KAA for storage in 80% ethanol Collect and store in 75–80% ethanol for later slide-mounting

Siphonaptera Trichoptera

Adults: can pin through mesonotum to right of midline, and spread wings, or collect and preserve in 80% ethanol. Larvae (with cases), KAA before storage in 80% ethanol

Lepidoptera

Adults: pin through centre of thorax and set. Larvae: KAA and store in 80–90% ethanol.

Hymenoptera

Larger adults; pin through mesothorax to right of midline; set as required, but commonly not done. Smaller adults on card rectangles or points: ants on card points with head to right. Otherwise store in 80% ethanol, possibly for later mounting on card or slides. Larvae in 80–90% ethanol

Specimen treatment | 41

baiting, sweep netting, benthic grabs, suction samples, etc.—again see Chapter 4) may be removed from the net or trap and also placed in a liquid preservative. Liquid preservation is the most common option, with the exceptions of (1) light-trap samples of moths and some other nocturnal insects, (2) other types of insects needed for specific purposes, or (3) in the case of specimens to be collected alive. An alternative to wet preservation is dry preservation. This is not suitable for all insect groups, or for larvae (Table 2.3), but dry preservation is essential for adult Lepidoptera, because scales are easily washed from the wings and body in liquid. Dry preservation is the conventional method for many insect groups, particularly for larger and harder insects. For some groups, a small series of dried and pinned or staged specimens (see below) can be accompanied by bulk material stored in alcohol. For liquid preservation of adult insects, 70–80% ethanol (ethyl alcohol) is the most common option, although more specialized fixatives may be needed when the material is to be used for genetic or histological studies. Larvae are better

Table 2.3 Major fixatives used for preservation of insect larvae before storage in alcohol Fixative

Formulation (volume units)

Uses

Carnoy’s liquid

Glacial acetic acid: 1 95% ethanol: 6 Chloroform: 3

Rapid penetration without hardening. As larval fixative, use for at least 2 hours, not more than 24 hours

KAA

Glacial acetic acid: 2 95% ethanol: 10 Kerosene (undyed): 1

Larvae; kill in fixative, then as for Carnoy’s.

FAA

Glacial acetic acid: 1 Distilled water: 20 Formalin: 5 95% ethanol: 25

Particularly useful for aquatic insects. Can store indefinitely, but insects may harden.

Pampel’s fluid

Glacial acetic acid: 2–4 95% ethanol: 15 Distilled water: 30 Formalin: 6

As for FAA

Lactic–alcohol

95% ethanol: 2 75% lactic acid: 1

Aphids and scale insects. Kill and store

42 | Taxonomy and curation of insects

treated in slightly more complex fashion, and may be fi xed and stored as in Table 2.3. At the time of initial retrieval from traps, insects should be transferred into fresh preservative, because the liquid used in traps may either be unsuitable for long-term preservation, or may have become overly dilute or dirty. Furthermore, loss of body fluid from insects can lead to dilution of alcohol preservative, so that it is good practice to replace the alcohol at least once more (after about a week) before permanent storage. At initial preservation, it is useful to remove (and preserve separately) any bycatch (Chapter 2.1), as well as of course removing contaminants such as vegetation and other debris. It is also very important that the catch from each single trap (or other defined sampling unit) at any one time be kept in a separate bottle, so that the unit remains intact. Also, it is essential at this time to put the data label, written in pencil or alcohol- and waterproof ink, inside the storage bottle. Individually captured larger, adult insects should be killed humanely in a killing bottle or jar as soon as possible after capture. Odonata, however, may be placed in a paper envelope and immersed in acetone overnight to partially preserve their colours, while also leaving them in a condition suitable for DNA studies. Larger-bodied orthopteroids should be kept dry after killing. A killing bottle is a wide-mouthed glass jar with tight-fitting screw top or cork plug lid, containing a layer (1–2 cm) of plaster of Paris (gypsum) on the bottom (Fig. 2.4). This is impregnated with an anaesthetic agent, and the insects confined in the jar are killed by exposure to the fumes. Two killing agents commonly used are ethyl acetate and potassium cyanide. A few drops of ethyl acetate on the plaster layer serves as a fumigant for at least several days before replenishment. Alternatively, crystals of potassium cyanide (care: extremely dangerous: check with your local authority whether you are allowed to use it) may be sealed under the plaster at the time of making the killing jar, preferably under a layer of vermiculite or similar expander. Potassium cyanide has the advantage that it is long lasting without replenishment, and (particularly in remote areas) obviates the need to carry liquid ethyl acetate in a separate, sometimes fragile, container. However, use of cyanide means that any discarded jars must be disposed of according to local regulations regarding disposal of toxic substances. The jar should anyway be wrapped securely in masking tape to guard against contamination from accidental breakage before disposal. Ethyl acetate has the advantage that the specimen is left very pliable, making it easy to set on a setting board (see below). Its disadvantage is that it can leave some specimens pink in colour, especially the legs of grasshoppers. Killing jars should be carried in a knapsack or belt-pack, not in the pocket where they are easily broken and could cause injury.

Specimen treatment | 43 (a)

(b)

Fig. 2.4 Designs for a killing jar for selected captured insects. (a) Plaster of Paris is set into the bottom of the jar and thoroughly dried. A teaspoonful of killing agent such as ethyl acetate is added to the plaster of Paris base, and before use some crumpled strips of tissue paper are placed in the bottom. (b) This is a permanently charged jar, with a layer of cyanide crystals, sealed in with a layer of vermiculite and then a layer of plaster of Paris left to dry. Tissue paper is placed in the bottom just before use. These jars must be labelled as containing poison, and also with a skull and cross-bones (not shown here for clarity). They must never be placed in the sun or in the pocket of clothing next to the body. Wrapping the jar in adhesive tape is also recommended, to prevent injury if the jar breaks. Shown here are narrowneck jars which are easy to handle and carry on a belt with a specially designed pocket. For insects with large wings (e.g. some butterflies) a wide-necked jar may be preferred. The cyanide jar is usually surrounded by some duct tape as an additional guard against breakage (not shown here for clarity).

While carrying out a survey (depending on the objective and target group/s), it is wise to carry several killing jars for different sites, for different samples, or for different insect groups. A jar used for killing Lepidoptera is not suitable for other insects, because it becomes contaminated with wing scales and is often difficult to clean subsequently. Insects should not be left in jars for longer than necessary, because addition of other, live, insects can lead to damage through trampling

44 | Taxonomy and curation of insects

and tumbling around. Placing one or two loosely crushed paper tissues in the jar can help prevent this damage. Some workers opt to transfer stunned insects to envelopes and place these in a larger, reservoir killing jar to liberate the normal killing jars for repeated use. Tiny or very delicate insects such as microlepidoptera can be collected individually into small glass vials and these, plugged loosely with cellulose wadding, can be placed into a killing jar; or a drop of ethyl acetate added directly to the plug before sealing these in a jar. As noted earlier, killing by freezing is needed for material to be used for genetic studies: in remote areas, a car fridge or polystyrene beer-cooler is a useful tool. Cellulose wadding is much preferred to cotton wool, as it does not snag on insect body parts. Depending on the freshness of the chemical killing jar, 1–3 hours is usually enough to ensure death, with the insects becoming comatose after much shorter periods. Specimens should then be transferred for storage or further preparation, such as pinning. Dry storage may entail papering (for butterflies, dragonflies, and other large winged insects), or layering for beetles and others. Papering involves placing the insects individually with wings folded back into triangular or other envelopes, preferably translucent, so that the contents are visible, and with data details written on the outside or enclosed with the specimen (Fig. 2.5). Where such envelopes are not available, or where conditions are very humid, envelopes can be made out of newspaper. Specimens can be kept like

1

a

b

2

b

3

4

Fig. 2.5 Folding procedure for making a temporary storage envelope for dead insect specimens. Note that there is only one specimen per envelope, and the envelope is labelled with all the appropriate data.

Specimen treatment | 45

this for long periods for future permanent mounting and storage (as long as they are protected from the ravages of pests). Envelopes can be stored in a rigid card or wooden box or cabinet drawer, with the outer container containing a small amount of naphthalene to deter damage from foraging insects such as ants, particularly at night in field camps. Layering entails using a plastic container, such as a sandwich box, in which a thick (~5 mm) layer of paper tissues is placed. A layer of the collected, dry insects is placed on this, a label is added, and the whole covered with more tissues without crushing. The process is repeated until the box is filled (Fig. 2.6), so that the insects are kept secure, uncrushed, and firmly enclosed so that they cannot get shaken around. If field sorting time is limited, it is useful to put like with like where feasible, so that all beetles or all flies are together in one box or in one layer. This shortens later sorting and possible damage of handling dry insects before they are prepared further. Specimens from different samples or sites should always be layered separately to avoid possible mixing. Also, they should not be left in such a sealed box for more than a few days without some sort of extra drying, otherwise mould will set in. In the tropics in particular, it is essential to add

Fig. 2.6 Loose insect specimens layered in a field storage box. Papered specimens (see Figure 2.5) can also be placed in such boxes. Specimens are kept like this until they can be prepared for permanent storage.

46 | Taxonomy and curation of insects

a mould suppressant such as thymol and a desiccant such as silica gel. Both these substances are normally available at well-stocked pharmacies.

2.7.1.2 Mounting Methods for mounting individual insects are summarized in Table 2.2. Other than slide-mounting, which is needed for critical identification of very small insects such as thrips and booklice, the main technique is direct pinning. This is often accompanied by some form of setting or spreading to display the various structures (such as wings) neatly and effectively, and staging. Direct pinning is the most frequent mode for many larger insects, using specially made entomological pins of stainless steel with lengths around 38 mm. Regular dressmaking pins are no good as they are normally too short and thick, of poorer quality steel, and corrode very quickly. Various thicknesses of entomological pin are available ranging from fine (0) to extra thick (5), with the (a)

Fig. 2.7a Position of the entomological pin, viewed from above, when preparing certain insect groups for permanent storage. Clockwise from top left: butterfly (or moth) (pin central), bug (pin slightly to right of midline), cricket (and grasshopper) (pin slightly to right of midline), beetle (pin through right elytron), bee and fly (pin to right of midline). (Specimens not to scale.)

Specimen treatment | 47 (b)

Fig. 2.7b Position of the entomological pin, viewed from the side, when preparing an insect (in this case a fly) for permanent storage. Only the specimen in the top left is correctly pinned; the other four are incorrectly pinned.

intermediate grade (3) being the most useful single category. Finer pins tend to bend, and thicker ones are needed only for the very largest insects. The position of the vertical pin differs for different insect groups. Lepidoptera, for example, are pinned through the midline of the thorax, but groups such as Orthoptera and Hemiptera are commonly pinned slightly to the side of the midline, to preserve midline features of possible taxonomic significance. Beetles are pinned through the right elytron, about one third distance from the midline and the same distance from the anterior margin. Figure 2.7 indicates some of these variations. Points to note for standardization (Fig. 2.7b) are: (1) that the pin is vertical, (2) that the position of the insects on the pins is standardized (to facilitate easy scanning under a microscope without need to continuously refocus) and leaving enough space above the insect for handling, and enough below for labels, and (3) that a suitable-sized pin is used. To standardize height of the pinned insects on the pins, a pinning stage is used, which can easily be made at home from a wooden or aluminium block (Figure 2.8).

48 | Taxonomy and curation of insects

Fig. 2.8 A pinning stage to ensure correct and consistent pinning heights of insect specimens. This can easily be made out of a small aluminium or wooden block. The shorter shafts of the stage can also be used for maintaining consistent heights of the data labels.

Insects can be pinned only when they are soft and pliable. When dead for more than a few hours, insects rapidly become dry and stiff. Attempts to pin them in this state may damage them. Note, however, that many insects undergo a period of rigor mortis, becoming temporarily stiff for a few hours and softening for a short period after about 10–24 hours. Subsequent stiffening is permanent. However, long-dead insects, for example in paper envelopes or layered in boxes, can be relaxed by exposing them to a humid environment for a day or so (for tough beetles, more rapid treatment by immersing them in hot water is sometimes an option). A suitable relaxing box can be made from a plastic sandwich box with a layer of clean sand (covered with paper to prevent contamination of insects) or tissues (without any soluble colour) that can be dampened with water to create a humidity chamber once the lid is closed (Fig. 2.9). Insects to be relaxed are placed in the chamber, on top of an additional layer of tissues to prevent direct wetting. They are then left overnight, by which time many will already be sufficiently softened to permit pinning. If not, test again after a further 6–8 hours, or after a second night. Specimens in this state should usually be handled with very soft forceps, which can be home-made from the steel straps used around wooden shipping cases (Fig. 2.10). If left unattended for longer periods, specimens may become mouldy. To prevent this, a small amount of thymol or other fungistat placed in the relaxing box will help to prevent this (NB: the use of chlorocresol, mentioned in some early texts, is not recommended as it is hazardous to health).

Specimen treatment | 49

Fig. 2.9 A relaxing box, made from a plastic sandwich box, layered at the bottom with damp white tissue. Specimens that have hardened during temporary storage are softened (and thus made ready for setting) by placing them for a day or two in such a relaxing box. A pinch of thymol prevents any build-up of mould.

Fig. 2.10 Very sensitive forceps, home-made out of packing case steel, for handling delicate insects, e.g. putting in and taking out of dragonflies from transparent envelopes. Also shown is a simple but very versatile and useful piece of field equipment—a good-quality 10ⴛ and 20ⴛ hand lens.

50 | Taxonomy and curation of insects

Very small insects can be pinned using tiny micropins (usually only 15 or 20 mm long, and of very fine stainless steel) for subsequent staging (see below). As above, the specimens must be thoroughly soft when pinned. Micropinning is a very delicate operation, usually undertaken with the aid of a binocular microscope. The specimen is supported with fine entomological forceps on a piece of lint-free cloth such as a handkerchief, or a block of expanded polystyrene or expanded polyethylene (plastozote) or other commercially available plastic foam used to line insect cabinet drawers, covered with fabric to prevent movement. Micropinned insects can be set (see below) on boards, or the micropins inserted into plastic foam in shallow plastic trays with the body in contact with the foam. Wings are then gently moved to the required position and, if necessary, supported by pins, and the insect left to dry over the next 3–7 days. Labels should be pinned alongside for later staging with the specimens. Many pinned insects, of all sizes, need some manipulation before they harden, to increase accessibility of taxonomic features and to augment their aesthetic appeal. To achieve this, they must be set, which means spreading their wings and legs (as appropriate) to expose as many of their significant morphological features as possible. Lepidoptera are usually set so that the wings of both sides of the body are extended horizontally and symmetrically, but many other groups of insects conventionally have wings extended on only one side. Where there is a long series of obviously the same species, not all specimens need be set in this way. This saves both time and storage space (see below). Besides, if needed in the future, additional specimens can always be relaxed and reset. Setting involves use of a setting board. This is a wooden board with cork or plastozote facing, with a central groove to accommodate the insect’s body, and a flat surface to support the wings and other appendages during drying (Figure 2.11) for 1–3 weeks, depending on size. The wings are carefully moved into position with fine mounted needles (setting needles), held in place by covering setting strips, ideally of translucent paper, and secured in position by pins, as shown Figure 2.11. Setting boards should be kept safe over the period of use. If left exposed, the insect specimens will readily be eaten by other insects or mice! Labels should accompany the insect on the setting board, and afterwards be attached to the specimen pin beneath the specimen itself. Partial setting, which involves simply parting the wings from the body, and moving other appendages as necessary for greater visibility, is another way of arranging insect specimens for ease of study. Staging refers to the insect being glued on to a card mount (or stage) and further supported by a normal entomological pin placed through the mount

Specimen treatment | 51

Fig. 2.11 Setting an insect, in this case two butterflies and a moth, to expose maximum area of the wings, while at the same time having some sort of naturalness. The butterfly at the top right still needs its wings moved up slightly, whereas the moth is its final position. The wings are spread with a setting needle, and held in position with paper setting strips and pins. Notice that the main pin through the body is exactly at 90° to the wings to ensure an orderly appearance. The data label, which may already be prepared in final form, accompanies the specimen. The set insects are left in this position, ideally for a few weeks, in a dry, protected, well-aerated place.

rather than through the insect. The insect can also be micropinned to the stage. Many small insects, too small or too hard to pin, are mounted directly on to card triangles or rectangles (Figure 2.12), a process termed carding or card-mounting. Micropinned insects are usually pinned near one end of short lengths (8–12 mm) of square cross-section foam, such as plastozote or similar compound plastic, and a regular pin inserted near the other end. Small beetles, ants, and parasitic wasps, for example, are normally card-mounted. They are stuck onto the tip of a small triangular or centre of a rectangular piece of card with water- or alcohol-soluble glue, so that specimens can be soaked off if need be. Do not use any glue that may blob or is not transparent, or any type of superglue that damages the specimens. Ants are normally placed across the tip of card triangles (Figure 2.12), but beetles are usually placed on a firmer, rectangular mount. If the material is to be deposited in a specific institutional collection, it is useful to seek specific curatorial advice on the procedures preferred by that

52 | Taxonomy and curation of insects

Fig. 2.12 Staged insect specimens may be mounted on the point of a card triangle, in the centre of a card or on the end of a strip for larger specimens, or micropinned onto a small block of plastozote. The data label sits below the specimen on the same pin. A wide card triangle is shown here, to leave space for any parts that accidentally break off or are deliberately removed to be water-glued to the card.

institution. All of the materials mentioned above are available from dealers in entomological supplies.

Specimen treatment | 53

2.7.1.3 Storage Adult insects with important, diagnostic soft parts and many larvae may be unsuitable for dry preservation and so are preserved in liquid preservative. Conversely, there are a few insects that should not be preserved in liquid as important diagnostic features, such as the white waxy covering of mealybugs, are lost. Insects in liquid preservatives (e.g. 75% ethanol) are stored in soda glass jars with tightly fitting lids (such as fruit-bottling jars with rubber seals) to stop evaporation (Figure 2.13). Individual insects can be stored in small soda glass vials and placed upside down in such jars. Regular inspection to top up preservative as needed is a continuing responsibility for the curator of the collection. Specimen data labels should be placed inside the jars with the specimens. Fire or other safety issues may dictate that the alcohol containers must be stored in special metal or fire-proof/fire-resistant cupboards, and may also impose other conditions, such as on the total volume of alcohol that may be stored. Pinned or staged insects are kept in wooden entomological storage boxes with tight-fitting lids, or in specially made insect cabinets, consisting of a series of glass-topped drawers (Figure 2.14). You should visit your local museum or other institutional collection to see how storage works in practice. The main purpose of storage is to protect the specimens from direct physical damage by exposure, from moisture, from excess light (that may cause fading of colours), and from damage by pests such as museum beetles (Anthrenus) which rapidly enter collections and eat their way through the precious specimens. The storage containers must therefore be virtually airtight. A small amount of a recommended, non-hazardous, solid insecticide, with an active ingredient such as d-allethrin (see, for example, the UK Health and Safety Executive website), can be placed inside the storage container and secured in position with a pin or a small cardboard box as necessary. This is not a permanent solution, and the storage container (box or drawer) must be inspected every few weeks to check that there are no invasive museum beetles. Should any mould appear on the specimens, it must be dealt with immediately by placing thymol crystals inside the storage box or drawer. When maintained well, a collection will last for many decades, and indeed, some museums have specimens that are well over 200 years old. It is worth spending the time to make up or buy an examination stage, which is a base with a swivelling arms and a rotating stage on which is glued a small piece of plastozote (Figure 2.15). This enables the pinned insect to be turned and viewed from various angles, especially under a binocular microscope, to assist with observation and identification. A cheap, simple alternative, although it

54 | Taxonomy and curation of insects

Fig. 2.13 Some insect specimens require wet storage, i.e. in a preservative such as 75% ethanol. Individual specimens, or small groups collected at the same time in exactly the same habitat, are placed in small vials, with their data labels, and plugged with cellulose wadding. The tubes are placed upside down (having expelled all the air so that they sink) in a large jar with a good seal, and the whole jar topped up with the same preservative. The vials should sit on a layer of cellulose wadding, and, if necessary, can be held in position by more wadding throughout the jar. Traditionally, wide-necked jars were used, but these tend to gradually lose the alcohol over time, so tightly sealed preservative jars with strong caps and robust seals are now favoured.

requires more handling of specimens, can be made from a blob of plasticine on a microscope slide or similar substrate that fits on the microscope stage. Again, most of the material needed for generating and curating insect collections are available from dealers in entomological supplies.

Specimen treatment | 55

Fig. 2.14 A well-curated permanent storage collection. This is a drawer from an insect cabinet, and has a very tight-fitting lid. Some d-allethrin is placed in the corner (to keep out museum beetles), along with a small tray of thymol (to keep out mould).

Fig. 2.15 An examination stage can be made by a local metal worker. The aim is to provide a stage on to which the insect is pinned for close examination, particularly under a binocular microscope, by turning through the vertical axis (with the lower knurled knob) and through the horizontal axis (with the upper knob).

56 | Taxonomy and curation of insects

If it becomes necessary to send dried pinned insects by post or courier to a museum or specialist, they must be protected carefully. Individual insects must be placed in a tough cardboard or wooden box with a base of firmly affi xed plastozote (do not use polystyrene as it does not grip the pin sufficiently and the specimens will shake free). They must be cross-pinned to prevent them coming loose or swivelling on the pin (Figure 2.16). The boxes containing the insects should be as small as practicable and sent within larger boxes containing expanded polyfoam chips, bubble wrap, or similar buffer to surround the insect box by 5–10 cm on all sides. Packages should be sent by the shortest direct route, and clearly marked ‘FRAGILE’, on the outside. For international travel, further precautions may be needed to avoid damage from opening in customs: any paperwork needed must be clearly available, and inspection of the contents without direct handling may be facilitated by covering the insect box before closing with a clear plastic wrap such as kitchen cling-film (Glad wrap, Saran wrap, or some similar product). Finally, it is worth emphasizing again that before embarking on preparation and storage of important insect survey material, it is really worth the time and effort to discuss your project with an experienced curator. Even limited consultation may save considerable work, avoid frustration, and avoid important errors. It might also be important to have access to a major collection during the identification phase of a study, and museum staff may be able to give advice on

Fig. 2.16 When pinned insect specimens are sent by post or courier, each entomological pin with its specimen must be placed firmly into the plastozote base. The specimens are then cross-pinned to hold them firmly in position. A layer of stretch-wrap kitchen film should be wrapped across the box before it is closed. Finally, the whole box is covered with bubble-wrap or immersed in polyfoam chips inside its outer postal box.

Summary | 57

relevant taxonomic literature during the planning stages. In turn, specialists may be able to provide most valuable aid as the work proceeds. Non-entomologists sometimes find it difficult to appreciate that, unlike workers on birds, reptiles, or mammals—all of whom may have a good working knowledge of, and be able to identify, most members of the group with which they are familiar—entomologists may specialize in only one order of insect, or even in only one family of a large order such as Coleoptera, Diptera or Hymenoptera. Usually they are still able to provide much general insight even when it is not their speciality, and where appropriate, recommend where you may be able to go to seek appropriate advice on other taxonomic groups.

2.8 Summary Before starting any field work, it is essential to plan well. In addition to designing a sampling protocol (Chapter 3), this includes how you are going to collect insects (see Chapter 4), and how you are going to preserve and store them (this chapter). In other words, planning involves thinking through all the practical steps before embarking on work in the field. This plan may be refined by doing some preliminary field work (a pilot study, see Chapter 3) to ascertain whether the plan is workable, and to more closely define what resources, both time and money, are going to be needed. Choosing the insect group for study depends very much on the precise aims of the project and how well the species are scientifically known in the geographical area in which you are going to work. We strongly recommend working at the species level, and not at higher taxonomic levels, such as family. This is because there is far more variation and sensitivity to environmental variables at the species level than at higher levels. It may be possible to actually identify sampled species, such as butterflies and dragonflies, even in tropical areas. But for many taxa it will only be possible for you to work on recognizable morphospecies, a collection of which is likely to be a close approximation of the real species assemblage. In geographical areas where some taxa are relatively well known and others less so, you may choose to do a study using both morphospecies and identified species. Where there are large numbers of samples to process, it may be possible to employ local parataxonomists, who can be trained to sort captured specimens into recognizable taxa, and in some cases, even identify the taxa. After sorting and thorough labelling of the specimens, it will be necessary to develop a voucher collection, which is effectively a permanent voucher collection to which you, and others after you, can refer. Such voucher collections, besides housing the actual selected specimens, may also contain

58 | Taxonomy and curation of insects

additional information such as digital images, so that there is a supplementary virtual voucher collection. It is critical for any study that the specimens are properly processed. While recommendations are given here for various insect taxa, we strongly advise that you contact the museum or other institution to whom you might send specimens for identification. They will advise on the exact method should be used for preserving, arranging and storing the collected material. For most taxa there are standardized techniques that should normally be adhered to, unless advised otherwise by a specialist. Finally, it is critical that your collection is carefully stored to enhance its long-term value. This involves, among other things, keeping out museum beetles and mould. All these considerations are part of the initial planning of the project, and should not be left until the specimens have already been collected.

3 Designing sampling protocols for insect conservation

3.1 Introduction Insects, by virtue of their variety, small size, and large number, do not readily lend themselves to being counted or their diversity quantified. They most often first have to be conned, cajoled, or otherwise extracted from their crevices and other hiding places using a selection of the techniques that are described in the next chapter. However, once the target group or taxon (for example grounddwelling or aquatic insects, such as ants or aquatic beetles) and most appropriate collection method(s) have been selected, several other questions arise. Ecologists (and entomologists in particular) are almost never able to obtain a complete measure of reality, and must rely on samples to ‘represent’ that reality. However, clearly we must do all that we can to ensure that the samples taken, and thus the outcome of the study, are as reliable (trustworthy) as possible. In other words, when sampling, we have to minimize bias and sampling error (Table 3.1). These arise from poor practical techniques (measurement or investigator error) or poor sample design (for example too few samples or poor distribution of samples as we will discuss in this chapter). Most of the following questions should be answered well before venturing into the great outdoors. In fact, asking and correctly answering the following set of questions will make the difference between the success (meeting the objective, obtaining reliable, information-rich results, successful publication of the research, and producing viable recommendations for management and conservation) or failure of your study. The important questions you should ask are: • • • •

When should I sample? Where should I sample (precisely rather than generally)? How often should I sample? How many samples should I take?

60 | Designing sampling protocols for insect conservation Table 3.1 Decisions to be made when designing a sampling protocol Question requiring a decision

Further discussion

What are the study objectives, key questions, and null and alternative hypotheses?

This chapter

What is the target taxon (taxa) or group (e.g. functional group)? Chapter 2 Is the taxonomic expertise available and accessible so that the species sampled can be identified?

Chapter 2

What are the variables and relational (environmental) variables to This chapter and be measured? Chapter 5 Where will the study be conducted and what will the extent of the study be?

This chapter

What is/are the most appropriate sampling techniques?

Chapter 4

What is the grain of the study?

This chapter

How many samples and replicates are needed (power analysis)?

This chapter

Where should the samples and replicates be placed across the study area (extent)?

This chapter

What is the duration, timing and frequency of sampling? Is a pilot study necessary?

This chapter This chapter

The answers to these questions will determine the sampling design of the study. These answers are dictated by four things: (1) the biology of the target group or taxon; (2) the sampling technique chosen (e.g. pitfall vs sweep-net) (see Chapter 4); (3) the structure and physiognomy of the habitat and landscape; and, importantly, (4) the objective of the study. Let’s start by considering the study objective.

3.2 Defining the conservation question How you go about sampling depends very much on why you intend to sample. In fact, the most important ingredients to the eventual success of any study are clear and precise statement of the study questions and objectives, and, thereafter, good planning (Figure 3.1). If you set out on your bicycle with no specific destination in mind, don’t be surprised where you land up! The same applies to research studies without clear objectives. Therefore, the more clearly and more detailed the objective is at the start of the study, the greater the likelihood that the outcome will achieve that objective. In conservation studies, managementrelated questions and information requirements form the priority objectives.

Defining the conservation question | 61 Start here

Observations Patterns in space and time

Models Explanations or theories

Hypothesis Predictions based on model

Null hypothesis Logical opposite to hypothesis

Experiment Critical test of null hypothesis

Retain Ho Refute hypothesis and model

Reject Ho Interpretation

Support hypothesis and model

Don't end here

Fig. 3.1 Generalized scheme of logical components of a research programme. HO represents the null hypothesis (from Underwood 1997).

3.2.1 Broad objectives in insect conservation There are generally only four broad objectives when sampling insects to answer ecological or conservation-related questions. These objectives are: 1) To assess (inventory) the species richness and species composition of an area. This will require a survey, and the outcome will be a species list for the area. For example, a nature reserve may want a butterfly list to supply to visitors of the reserve. In a survey, one will often use as many

62 | Designing sampling protocols for insect conservation

different sampling techniques as is practical to obtain as complete a list of species as possible. A survey may also be more targeted and merely involve confirming the presence or, with more difficulty, the absence of a species across a series of sites, or to sample specimens for genetic analysis. 2) To understand the habitat or environmental associations (or habitat requirements) of one or more species. This objective requires more than a survey, and a range of habitat types (e.g. forest, grassland) or environmental states (e.g. early and late succession habitats, polluted and unpolluted) will have to be sampled (and as we will see later, adequately replicated). The outcome of this objective will be a species list (and usually associated abundance or density values) for each habitat type or environmental state. Here, the same sampling techniques and sampling effort must be applied to each habitat type or environmental state to ensure comparability of results between them. 3) To understand the impact of environmental change on one or more species. One or both of the following approaches (sensu Hurlbert 1984) can be adopted here. First, the study may be mensurate, i.e. quantifying the environment in its natural or current state (also called a mensurate experiment). Alternatively, the approach may be manipulative, where the environment is deliberately manipulated, or altered, in a desired way. For example, a study in which you sieve the tree bark for saproxylic beetles in existing stands of logged and unlogged boreal forest would be mensurate. However, if you were to negotiate in advance with the forest managers to log pre-selected forest stands at different intensities (say cleared stands, stands with half the trees removed, and other stands with no logging) and then sample these, the study would be manipulative or experimental. Insect ecophysiology studies are often experimental, where you would, for example, measure the developmental rate of a lepidopteran species at a range of temperatures in the laboratory. Insect conservation studies are commonly mensurate, although where possible manipulative studies are often more convincing (indeed arguments based on a combination of the two approaches are particularly powerful, see Dunne et al. 2004). For many rare insect species, however, when only one or a few populations are known, manipulation may be too risky. 4) When dealing with rare, threatened, or ecologically important species, the objective may be to monitor changes in the population size (or structure of a community when more than a single species is involved) over time (e.g. forming part of an adaptive management strategy for a rare species). One may also be interested in assessing how the population size

Defining the conservation question | 63

of the species responds to changing climate, wet vs dry seasons, different levels of harvesting or other disturbances, perhaps over long periods. These sorts of studies involve collecting time series information (measurements collected, usually at regular intervals, over time). They are also sometimes called longitudinal studies.

3.2.2 Refining the objective Establishing which of the above broad objectives is the case (and a research programme may well encompass all four) is only the first step. Objectives can and should be described in as much detail as possible. In most studies, the broad objective becomes a lot more complex and detailed once you start thinking about it. Take, for example, the following objective: to quantify the effect of over-browsing by elephants on insect diversity. First, quantifying ‘insect diversity’ in an African savanna (or most other habitats) is no small task, and you would probably want to select one or more groups of insects to work with, say dung beetles or herbivorous insects. This decision may be based on previous studies showing that, for example, dung beetles are particularly sensitive to browsing or changes in habitat structure. Alternatively, it may be based on logistic grounds, e.g. dung beetles are comparatively easy to sample in a quantifiable fashion, and the dung beetle fauna for the area of interest is well known. When making this decision, it is important to make sure that you have a good understanding of the biology and behaviour of the taxon (or taxa) concerned, and that you have, or will readily be able to attain, the taxonomic expertise to identify the samples that are collected (see Chapter 2). The taxon (taxa) chosen will influence the decision on when to sample, how long to sample for and at what intervals, and which sampling technique to use (Chapter 4). The ‘area of interest’ also needs to be defined and may be, for example, a country, region, biome, habitat, or series of nature reserves. Next, are we interested in impacts that are short-term (a few days to one month after browsing) or longer-term (increasing loss of forest canopy or decline in tree densities over time), or impacts within or across seasons? The answer to all of these questions will determine the way in which the study is designed (Table 3.1). As a result, time and careful thinking must be dedicated to clearly defining the study objective and considering the best ways to approach it. Such planning and refinement is a sound investment for the future of your work. In some cases, the precise objective of the study may be imposed by contractual demands, or the recommendations of action plans. You should carefully assess such objectives, and any constraints on the sampling design set by such objectives, before committing to a study. Sometimes contracts and action plans

64 | Designing sampling protocols for insect conservation

are not compiled by ecologists, and will need further development or modification to ensure that the desired outcome is achieved.

3.2.3 Hypothesis formulation It is most useful if one has a formal hypothesis in mind (mindless data-gathering seldom results in any concrete achievement) (Figure 3.1). For example, previous research may have shown that over-grazing in savanna causes a decline in the species richness of understorey herbivores. The hypothesis would then be that over-browsing by elephants results in a decline in canopy and middlestorey insect herbivores. The null hypothesis (or hypothesis of ‘no effect’) in this case, would be that elephant over-browsing results in no change to the insect herbivores (all good hypotheses have a null and alternative form; see ShraderFrechette and McCoy 1992; Figure 3.1). In another example, an increase in elephant activity will result in increased dung concentration in the area, which may result in an increase in dung beetle species richness in the short term (note that to test this hypothesis one would want to test whether dung concentration is indeed higher in over-browsed areas, as well as whether dung beetle species richness is higher). Browsing has also been shown to increase insect gall densities in some systems by increasing the vigour of new shoot growth (to test this hypothesis one would need to measure, for example, both new shoot density as well as gall abundance). This example relates to a well-known formal hypothesis in insect ecology, namely the plant-vigour hypothesis (that insect herbivory is higher on high-quality hosts, Price 1991a), and an excellent entomological example of the importance of hypotheses in the development of ecological theory (Price 1991b). These examples also illustrate that in most instances data on both direct variables (also called dependent variables) as well as relational variables (also called independent variables) must be collected. Relational variables usually include weather, climate or soil parameters, measures of habitat quality (plant diversity of cover), or as in the example above, shoot density or elephant browsing intensity (see Chapter 5 for further discussion of relational variables). One form of relational data that will be discussed in this chapter is spatial referencing (see section 3.3.5). The process of defi ning the objective and formulating hypotheses helps to identify the variables that must be measured, as well as the timing, duration, spatial position, and spatial extent of the study (sampling or study design). Therefore, the more clearly the objectives are described (most formally as hypotheses), the greater the success of the study will be.

Designing a sampling protocol | 65 Table 3.2 Possible outcomes of statistical hypothesis tests (from Scheiner and Gurevitch 2001) Reality (‘truth’)

Decision: based on study results Do not reject null hypothesis Reject null hypothesis

Null hypothesis is true Correct (1 – α) Null hypothesis is false Type II error ( β = the probability of making a type II error)

Type 1 error (α = the probability of making a type I error) Correct (1 – β; also considered to be the statistical power of the test)

Schrader-Frechette and McCoy (1992) discuss the importance of hypothesis formulation in ecology. They also point out that in applied sciences, such as conservation, it is often preferable to incorrectly reject a true null hypothesis (type I error, see Table 3.2) than not to reject a false null hypothesis (type II error, see Table 3.2). For example, take the following hypothesis: an upstream industrial development will change the habitat quality for a rare mayfly species (the null hypothesis would be that the development would not change the habitat quality). It would be better (an ethically based decision) to make a type I error than a type II error during the analysis stage of the study. In this case making a type I error (rejecting the null hypothesis) is equivalent to erring on the side of caution. Th is is preferable in this case to making a type II error (concluding that the development does not affect habitat quality) and being responsible for the extinction of a mayfly species! If the hypotheses in a study are not clearly formulated, it is difficult to evaluate these sorts of implications and consequences of the outcome of a study. Note that type I errors are not always preferable, and in fact scientists generally place more emphasis on avoiding type I errors, or prefer to err on the side of ignorance (SchraderFrechette and McCoy 1992).

3.3 Designing a sampling protocol Once the objective of the study is known and questions and hypotheses have been clearly formulated, a sampling protocol must be designed that will allow the questions to be answered and the hypotheses to be tested. The principles below are broadly applicable, and relevant, for example, to studies quantifying the spatial genetics and population structure of a species, to those quantifying species diversity, or manipulating the environment to examine disturbance effects on

66 | Designing sampling protocols for insect conservation Box 3.1 Elements of sample design The basic elements of sample design (sample grain, sample number, and sample extent) are illustrated below. When these elements are combined they determine the sampling coverage and sampling intensity of a study. In A, sampling coverage is 100% using either the smaller or larger of the two grain examples shown. Sampling intensity is, however, greater with the smaller of the two grains. The extent divided by the grain (both measured as area) is called the scope of the study. Progressing from B to C to D is equivalent to a decline in coverage and intensity (as extent increases, while sample number and grain remain constant). Different combinations of grain, sample number, and extent therefore alter the coverage and intensity, or scope, of a study. (A) Grain (area of smallest sample unit, sample number (n) = 16 in this case) Alternative grain example, n = 4 in this case Extent (size of area within which study is conducted) (B)

(C)

(D)

diversity. The three basic elements of sample design are sample grain, sample extent and sample number (Box 3.1).

3.3.1 Sample grain Sample grain is the physical size of the minimum resolvable sample unit (Box 3.1). The grain of the study is usually determined by both the study objective and the appropriate sampling technique or method. There are two ways in which the grain of a study may be determined. Grain could be identical to the size of the physical sample unit used, such as a 1 m2 botanical sampling grid, length of transect, or area covered by one or more pitfall traps. Alternatively, grain may be the

Designing a sampling protocol | 67

area of the unit to which occupancy is extrapolated, e.g. the presence or absence of species in geographic grid squares (representation-based sample units) (Gaston 1994). The latter represents a coarser sampling resolution (lower sampling intensity) than the former. Often, however, more than one trap or sampling event is used to make up a single sample (i.e. the grain of the study is made up of more than a single physical sample unit). For example, a grid of 10 pitfall traps (say in a 2 × 5 array) may be used as a single sample (usually to obtain enough individual insects to represent the insect diversity at that point). The data for these 10 pitfalls are pooled during analysis, and the effective grain is then the area covered by the grid of 10 pitfall traps, and not the area covered by a single pitfall. In other cases, the area covered by a single pitfall could form the grain of the study. The sample grain is equivalent to the sampling unit when the data are entered into a data matrix. Ludwig and Reynolds (1988) give an excellent overview, which is well worth consulting. Most importantly, although you may pool the data during analysis, never pool the actual individual samples themselves. If you do, it will not be possible to calculate, for example, between-pitfall variation in catch. You can always amalgamate data on a spreadsheet, but you can never separate the individuals from carelessly pooled samples.

3.3.2 Sample extent Sample extent is the physical size of the area within which the study is conducted, or the area of inference. It is important to decide what the extent is, because this is the area to which the result of the study may be extrapolated. Your sampling extent may, for example, be a game reserve, or a woodland, or a series of mountain valleys. Often, the study extent is predetermined, either by contract or by biology. Many insects of conservation concern are known only from a single small area, sometimes little more than a hectare or a few such areas (e.g. the threatened Brenton blue and Eltham copper butterflies). For example, if the question has to do with a particular wetland or farming region, then the area of the wetland or of the farming region forms the extent of the study. Once this context is known and specified, then a decision can be made about where the samples will be placed within and across the area, and how many samples are needed to provide a representative estimate for the area. If this process is completed correctly, then the results of the study may be inferred to hold for the full extent of the study area (although extrapolation beyond the focal taxon or habitat would of course not be valid).

68 | Designing sampling protocols for insect conservation

3.3.3 Sample number The sample number is the number of individual units with the same grain used in the study (e.g. if the area of one pitfall is the grain of the study, then the number of pitfalls provides the sample number; if an array of 10 pitfalls is the study grain, then the number of such arrays is the sample number for the study). The sample number in a study is usually a compromise between what is logistically possible (this sets a maximum limit—more sampling means more sorting time and greater cost) and the minimum number of samples needed to provide a precise estimate of the variable of interest (where the variable may be, for example, the number of butterfly species in the area, or the number of individual host plants) (Table 3.3). As a rule of thumb, three is the minimum number of samples needed for statistical estimation. However, generally speaking, the more samples taken, the more precise the estimate and thus many more than three samples are usually a good idea (10 replicates is comparably safe). Also, the more heterogeneous the environment in which you are sampling, the more samples will be needed for a precise estimate, in other words more samples will be needed to capture the variability in that environment (Table 3.3). In insect ecology and conservation, we are often interested in knowing how many insect species there are in an area or habitat. A small sample size is likely to fail to record the presence of some rare species within the sample area (McArdle 1990). A small sample size will also underestimate the real level of occupancy of some species and overestimate that of others. Overall, an increase in sample number tends to bring about an increase in the number of rare species found (McGeoch and Gaston 2002), because the generally large number of rare species in assemblages will only be captured with greater sample effort (see taxon sampling curves in Chapter 9). Sample size thus has an effect on the precision of estimates (e.g. means, slopes of relationships) obtained (Table 3.3). However, it also affects the likelihood of detecting statistically significant differences between groups (or treatments), or significant relationships between variables, for a specified significance level (α = 0.05 or α = 0.01; note that α here is the significance level and is not the same as that in Table 3.2). For example, the greater the variability associated with a particular variable, the more samples are required to detect statistically significant differences between groups, or relationships between variables (if such differences or relationships indeed exist). Therefore, if the sample size in a study is not large enough, the wrong decision may be made about the null hypothesis (Table 3.2). Power analysis is a formal process by which the implications of using different sample sizes in a study may be explored (Box 3.2).

Designing a sampling protocol | 69 Table 3.3 Definitions of bias, precision and accuracy (from Hellmann and Fowler 1999; Walther and Moore 2005) Definition

Implication for research result

Bias

Bias leads to underestimation or The difference between the expected value, sample overestimation of the true value mean, or test result, and the true (usually unknown) value. Non-random or directed error

Precision

Lack of precision leads to instability The statistical variance associated with the sample in the value of repeated measurements of the same object or variable data, as a consequence of measurement error and inherent variation in the variable of interest. Also referred to as ‘repeatability’

Accuracy

The less biased and more precise an estimate (such as the mean of a sample or a species richness estimate), the more accurate the estimate

Low accuracy leads to potentially misleading research results

Box 3.2 Statistical power and power analysis (a rudimentary knowledge of statistics is assumed here) Deciding on the number of replicates (number of independent samples, or sample size) in a study is an important component of designing a sampling protocol. Statistical power analysis is a tool that is used to make decisions about sample size (amongst others) during this process. It is used to explore how components of the prospective study will affect the likelihood of the study achieving its objectives. The power of a test is the chance that the test will detect a departure from the null hypothesis (1 – β, Table 3.2). The statistical power of a test is the probability of correctly rejecting the null hypothesis given that the alternative hypothesis is true (Scheiner and Gurevitch 2001) (see Table 3.2). The statistical power of a test increases with an: • Increase in sample size (n) • Increase in significance level (α) • Decrease in variance (σ2) • Increase in effect size (ES)

70 | Designing sampling protocols for insect conservation Power ~ (ESα√n)/σ Power is proportionately related to the relevant parameters, because the way in which power is calculated varies depending on the statistical test being used (Quinn and Keogh 2002). Effect size (ES) In power analysis, efffect size is the difference between the null hypothesis and a specific alternative hypothesis (Scheiner and Gurevitch 2001). For example, if the null hypothesis is that there is no difference in insect species richness between organic (So) and traditional (St) tomato fields (So – St = 0) and the alternative hypothesis is that insect species richness is 25% higher in organic than traditional tomato fields (where S, the total species richness = 100), then effect size is 25 – 0, i.e. ES = 25 species. Deciding on effect size can be difficult, but should be driven by what is considered to be a biologically meaningful difference in the situation of interest. Effect size is specified by the researcher and is a ‘hypothetical value that is determined by the null and alternative hypotheses’ (Scheiner and Gurevitch 2001). Nonetheless, statistical tests are more likely to detect large effects. Power analysis When used during the sampling design phase of the study, power analysis is used to determine: • The number of replicates or samples needed to ensure a specific level of power for tests of the null hypothesis, given a set effect size, α, and variance. • The power that a particular test will have when the maximum number of replicates logistically possible is used, with set effect size, α, and variance. • The minimum effect size that can be detected, given a target level of power, α, variance, and sample size (Scheiner and Gurevitch 2001). When conducting power analysis, by convention α is set at P = 0.05 (although the P-level may be selected as, for example 0.1 or 0.01; see discussion in Underwood 1997). A power of 80% is also often used by convention (1 – β = 0.8) (Quinn and Keough 2002). Estimates of the variance may be obtained by similar or related studies published in the scientific literature, or from a pilot study. A variety of statistical software that conducts power analysis is available, e.g. GPOWER and PASS (see review by Thomas and Krebs 1997) (summarized from Scheiner and Gurevitch 2001; see also Underwood 1997, Quinn and Keough 2002)

3.3.4 Sample independence It now becomes important to make the distinction between sample and replicate. When two samples are independent of each other then they may be considered to be replicates. If two samples are not independent of each other, they are not true replicates. The concept of sample independence is an extremely important one in sampling design, and relates to the problem of pseudoreplication, i.e. inadequate or incorrect replication (Figure 3.2). When a study is pseudoreplicated,

Designing a sampling protocol | 71

Replication

One quadrat (= sample grain) N.B. alternatively, it could be all four quadrats

Outer edge of forest reserve (= sampling extent)

Pseudoreplication

= Forest

= Selectively logged

Fig. 3.2 Example of a well-replicated (above) and pseudoreplicated (below) study. The objective of the study is to examine the effect of logging on forest insects. There are two treatments, i.e. unlogged (the control, shaded) and selectively logged (unshaded). The area of interest or inference, i.e. the sample extent, is the whole forest reserve. In the well-replicated example, traps are placed across several logged and unlogged blocks of forest. In the pseudoreplicated example, trapping effort is concentrated in only a single block of logged and unlogged forest. Although many more traps are placed in these single blocks, the study is not replicated at the correct level, i.e. at the level of forest treatment (logged and unlogged). The above example provides much better inference for the effects of selective logging across the whole reserve (sampling extent). The smallest unit, say a single quadrat or trap, is the sample grain or sampling unit. Note that in ecological and conservation studies it is not always possible to have a strictly randomized block (as is the case here), as it is with planned agricultural trials.

72 | Designing sampling protocols for insect conservation

the significance of statistical tests will be incorrectly interpreted, because the true value of α (see Box 3.3) is unknown (Hurlbert 1984). This means that the study objective will not be met and the investment in it wasted. It is clear that pseudoreplication should be avoided at all costs! The total number of replicates needed in a study is also dependent on the number of treatments (and may also be assessed using power analysis, Box 3.2). Although the word treatment is more formally used in manipulative experiments, it is equally useful in many mensurate (non-manipulative) field studies. Treatment may be, for example, different forest successional stages, elevational bands, disturbed vs undisturbed habitat (all mensurate), or different temperature manipulations, genetically modified vs unmodified crop varieties, or different larval diets (all manipulative). A certain number of replicates will be needed for every treatment, and when treatments are not replicated this also constitutes the ‘crime’ of pseudoreplication (Box 3.3). Box 3.3 Pseudoreplication Pseudoreplication is the use of statistics to test for treatment effects in experiments (manipulative or mensurate; see text) where: • replicates are not independent of each other (see also section 3.4.5), or • treatments are not replicated (Hurlbert 1984). Replicates are not independent of each other when those replicates influence each another in some way, or are influenced in the same way by an external factor that is not of primary interest to the research question. To avoid non-independence, replicates in mensurate studies should be isolated from each other in space or time (see also section 3.3.5, Figure 3.3). Example Consider each of the sample units in Figure 3.3 to represent a pitfall trap. Each catch of ground-dwelling organisms is a sample. Samples from individual pitfalls that are isolated from each other by a physical or geographic barrier are independent of each other, and are thus true replicates. This is all very well when useful or meaningful physical barriers exist in the study area. Far more often, individual pitfall catches (or other sample units) are taken to be independent of each other when the pitfalls are separated by a certain minimum distance (see section 3.3.5). Deciding what this ‘minimum distance’ is, is a tricky business. The decision is usually based on some understanding of the dispersal distances or capabilities of the target taxon. The idea is that for each pitfall to be independent, it should draw on a pool of fauna that is separate from, unique or independent of the faunal pools being drawn on by other pitfall traps in the same study. A general rule of thumb is that the farther apart two samples are from each other the more likely they are to be independent. Independence can be quantified by measuring the spatial autocorrelation of variables across the study area (3.3.5).

Designing a sampling protocol | 73 Treatments are not replicated when there is only a single replicate per treatment (per successional stage, elevational band, etc.). From the example below, it is clear that replication is possible at several levels. The appropriate treatment level at which replication must be used to avoid pseudoreplication is dependent on the question the research aims to address. Replication is usually also used at other levels to capture natural variability in the system and improve the precision of estimates (such as the mean) (see Table 3.3). Example Question

Pseudoreplication

Replication

Is there a difference in the insect diversity on trees growing … … at the edge vs the centre of Forest X? (treatment = tree position with two levels, i.e. edge and centre)

One tree is sampled at the edge of Forest X and one in the centre.

Several (independent) trees are sampled at both the edge and at the centre of Forest X

… at the edge vs the centre of forest patches in Nature Reserve Y? (treatments = forest patches and tree position)

One forest patch is sampled in Nature Reserve Y (with several trees at the edge and in the centre of the single patch)

Several (independent) forest patches are sampled in Nature Reserve Y (including several trees at the edge and in the centre of each patch)

… at the edge vs the centre of forest patches in nature reserves in Country Z? (treatments = nature reserves, forest patches and tree position)

One nature reserve is sampled in Country Z (including several forest patches in the reserve and several trees at the edge and in the centre of each patch)

Several (independent) nature reserves are sampled in Country Z (including several forest patches in each reserve and several trees at the edge and in the centre of each patch)

3.3.5 Spatial autocorrelation Two samples are spatially autocorrelated when they are more similar to each other than expected by chance (Legendre and Legendre 1998). This means that the samples are spatially non-independent (samples may also be temporally autocorrelated when collecting is repeated over time). Measuring the spatial autocorrelation amongst a spatial series of samples in a study is thus a way of quantifying the non-independence discussed in Box 3.3 and section 3.3.4, i.e.

74 | Designing sampling protocols for insect conservation

Isolated

Interspersed and not isolated

Interspersed

Isolated and interspersed

Isolated and not interspersed

Fig. 3.3 Illustration of the concepts of isolation and interspersion of replicates for one or two treatments (black and white). Here the ‘isolation’ is used to achieve independence of samples. This isolation may be in the form of a geographic barrier to dispersal of individuals, or merely distance (the farther away two samples are from each other, the more ‘independent’ they are likely to be). Top row from left to right: Isolated (1 treatment, 3 replicates); interspersed (2 treatments, 4 replicates); isolated and interspersed (2 treatments, 3 replicates); interspersed and not isolated (2 treatments, no replicates); isolated and not interspersed (2 treatments, 1 replicate).

if two samples are spatially autocorrelated, then they are not independent and cannot be considered replicates. If spatially autocorrelated samples are treated as replicates then their inclusion as such in statistical analysis unfortunately qualifies as pseudoreplication. One of the most common measures of spatial autocorrelation is Moran’s I, the values of which are usually presented in the form of a correlogram (Figure 3.4). Moran’s I lies approximately between –1.0 and +1.0, and values significantly greater than zero (or not significantly different from zero) demonstrate the presence of significant spatial autocorrelation. Values that are significantly less than (or no different from) zero identify distances over which values of the variable are significantly less similar (or no more similar) than expected by chance (and thus independent). Spatial or geographic coordinates (x and y or latitude and longitude) must be taken for each sample, forming a relational variable, in order to quantify the spatial autocorrelation across samples. As a rule of thumb, a minimum of 32 samples with unique spatial positions is needed to quantify spatial autocorrelation.

Designing a sampling protocol | 75 0.6 0.6 0.6

Moran's I

0.0 –0.2 –0.4 –0.6 –0.8 –1.0

0.9 1.8 2.6 3.5 4.4 5.3 6.2 7.0 7.9 8.8 9.7 10.6 Distance (km)

Fig. 3.4 Correlogram of the density (number of galls per plant) of a leaf-galling sawfly species (overall correlogram significance, P < 0.0001). Closed symbols (Moran’s I values ± s.d.) in the correlogram are significant at P < 0.05 (after progressive Bonferroni correction (α = 0.05)). This correlogram shows that gall density on plants within approximately 3 km of each other is more similar than you would expect by chance. In words, gall density measured from plants within 3 km of each other cannot be considered independent. (From McGeoch and Price 2004).

The measurement of spatial autocorrelation is used in another very interesting way in studies of insect populations or assemblages and relational variables that encompass sampling at several points in space. Here, spatial autocorrelation is used to estimate the distance over which values of a particular variable (e.g. species abundance, species richness, temperature, soil nutrient content) are significantly more similar to each other than expected by chance (or the distance at which they become significantly more different from each other than expected by chance). This provides a measure of patch size (or gap size), and can be used to generate hypotheses about the processes structuring the population or assemblage. For example, leaf gall abundance (expressed as gall density) in Figure 3.4 is positively spatially autocorrelated over distances of 2.6–3.5 km (an estimate of leaf gall patch size). Because sampling took place in a linear fashion along a river drainage system, this result shows that, on average, stretches of river drainage approximately 3 km long encompass similar densities of leaf galling sawflies (McGeoch and Price 2004). Measuring the spatial autocorrelation of variables is thus useful both for quantifying areas across which samples are non-independent, as well as for quantifying the spatial structure of populations

76 | Designing sampling protocols for insect conservation Random

Systematic

Stratified

Stratified-random

Direction of environmental gradient

Fig. 3.5 Alternative arrangements for sample placement across the extent of a study area. When there is a dominant environmental gradient in the study area, the extent is stratified perpendicular to the direction of the environmental gradient.

and assemblages. Spatial analysis in ecology has become a field in its own right (see Fortin and Dale 2005), and spatial position of sites and samples must now always be considered in studies conducted over space (Legendre 1993). Therefore, always record the spatial position of each sample in the study and remember that you may have to quantify the spatial autocorrelation of variables across the study area to: • identify the distance at which two samples (or values of a variable) become independent of each other • justify or demonstrate that samples are spatially independent of each other, or • describe the spatial structure of a variable across the extent of the study.

3.3.6 Sample placement Where you put your traps, or where you collect insect individuals (i.e. the position of each sample in the study area) is sample placement, and several terms are used to describe the way in which this is done. These terms (random, stratified, systematic, and also interspersed) all have to do with the way in which samples are arranged within the extent of the study (Figure 3.5). Randomization is the random assignment of samples to particular treatments, or the random arrangement of replicates within the extent of the study. Randomization prevents

Designing a sampling protocol | 77 A

B

C

D

E

F

Fig. 3.6 Randomized and systematic arrangement of replicates in a mensurate (A–C) and manipulative/experimental (D–F) study. A–C. Placement and orientation of 100 m sweep net transects in a patch of grassland in a mensurate study. D–F. Experiment to assess the effect of shading (100%, 50%, and no shade) on grassland insect diversity (manipulative study). A, transects positioned and orientated randomly; B, transects positioned and orientated systematically; C, transects positioned systematically and orientated randomly; D, treatments (full shade, half shade, and unshaded) assigned randomly to grassland plots (inadequate placement); E, treatments also assigned randomly (improvement on D); and F, treatments assigned systematically to grassland plots (best option).

the investigator from making biased decisions (deliberately or unwittingly) about where the replicates should be placed, or which replicates to assign to a particular treatment (Hurlbert 1984) (Table 3.2). Strangely, the potential downfall of randomization is that sample positions and assignments may by chance appear completely non-random. For example, in Figure 3.6A, it appears as though the south-west side of the patch was avoided when transects were placed, but this was in fact a random assignment of transects using randomly drawn x–y coordinates to place the centre of each transect (note that samples within a transect are by default not independent of each other). In this case, it may be better to place transects systematically (equal distances apart, where that distance is determined by the extent of the patch and the sample size) over the entire patch (Figures 3.5 and 3.6 B–C). If there is a known environmental gradient across the study area (e.g. shade, slope, temperature, soil quality) one may want to stratify the area, i.e. divide it into blocks perpendicular to the direction of the environmental gradient. Samples are then placed in each

78 | Designing sampling protocols for insect conservation

Ecotone

16 m

8

4

21

12

4

8

16 m

Sampling points at increasing distance from ecotone on both sides of the edge Habitat 1

Habitat 2

Fig. 3.7 Illustration of log2 sample (or trap) placement approach to sampling across habitat edges or ecotones. The minimum distance used depends on the habitat structure, and realistically usually starts at 2 m (excluding the 1.0 m sampling point).

block (Figure 3.5). In Figure 3.6C, transect centres are systematically placed and compass directions randomly assigned to each transect. Placing transects systematically across the patch will achieve good coverage (and thus a result that is representative of the patch), whereas the random orientation of transects will avoid any bias as a result of unknown environmental gradients across the patch (Figure 3.6C). Therefore, these sample placement approaches may be used together to achieve unbiased, representative results for the extent sampled. Generally, where there is more than a single treatment (e.g. habitat type or disturbance level), the replicates of each treatment should also be interspersed (Hurlbert 1984) (Figure 3.3). Interspersion, along with isolation by geographic barrier or distance, is another way of avoiding non-independence (Figure 3.6). As fragmentation is such a major issue in contemporary conservation, studies often involve close focus on the edge, interface, or ecotone between two vegetation types or land uses. The study may, for example, be about contrast (Wiens et al. 1993) between two habitat types, or penetration of the two habitat types by various species, i.e. landscape variegation (Ingham and Samways 1996). The point is that the focus is primarily on the edge, and secondarily on increasing distances away from the edge. The sampling is thus more intensive at the edge than away from it. Also, fragmentation studies to date have shown that edge effects often occur over short distances away from the edge (Ries et al. 2004). For insects, a suitable scaling for sample placement is m log2, starting at the edge (Figure 3.7). Sampling is then done at 0, 1, 2, 4, 8, 16, 32, 64, …, n metres from the edge, as far into the habitat adjacent to the edge as necessary. In practice, 1.0 m

Designing a sampling protocol | 79

may be simply too small a distance, so sampling is usually done at 0 m, 2 , 4 m, etc. A similar approach can be taken when studying road-zone effects, with sampling done in a log2 fashion from the edge of the road, path, or indeed any other linear corridor into adjacent habitat.

3.3.7 Sampling coverage and intensity Sample unit size (grain) and sample area (extent) define the lower and upper spatial limits of any study of organisms sampled across space (Wiens 1989). These basic elements are combined to produce different sampling coverages and sampling intensities (Box 3.1). Grain and extent, in addition to the number of samples taken, are often interdependent and together may be expressed as either coverage or sampling intensity. The latter are seldom considered components of sampling design, but do have an impact on the outcome and interpretation of the results. For example, the larger the grain relative to the sample extent, the more widespread species and fewer localized species there are likely to be sampled when there is complete coverage of the sample extent (McGeoch and Gaston 2002). As sample extent increases, the sampling intensity of studies tends to decrease as a result of logistical constraints (Wiens 1989; Gaston 1996). Although grain and sample number may be standardized as sample extent changes, the resultant change in sampling intensity may alter the way in which numbers of rare and common species are interpreted to change. A reduction in sampling intensity is likely to result in the loss of fine-scale details and an increase in differences between sample units (Wiens 1989). Low-intensity studies may therefore have apparently high predictability and patterns emerging from these will be far more readily generalized than from studies conducted at higher sampling intensities (Wiens 1989). At low sampling intensity, distances between samples may also be too great (or the grain too coarse) to detect smallerscale environmental gradients.

3.3.8 Controls The concept of control comes from manipulative experiments where one set of replicates is left ‘untreated’ or not manipulated. This control set of replicates serves as the baseline against which other treatments are compared (e.g. the unshaded plots in Figure 3.4). However, controls are also relevant to mensurate studies. For example, sampling the aquatic invertebrates in a river before a pollution event serves as the untreated control, or baseline condition, for aquatic invertebrate diversity in the absence of pollution. Similarly, simultaneously sampling several rivers, some of which are polluted and others not, provides treatment and control cases. Another form of control may be comparatively

80 | Designing sampling protocols for insect conservation

undisturbed or ‘natural’ habitat, in comparison with various states of anthropogenic disturbance. Procedural controls are used in manipulative experiments, where the effect of the application of the treatment must be separated from the treatment effect itself. For example, the procedural effect of the application of a liquid insecticide may be the moisture effect on insect diversity, rather than the insecticidal content of the liquid. In this case, the procedural control would involve the application of the same volume of liquid as in the insecticide treatment, but without the active ingredient. Controls are also often necessary to control for changes over time in a system (for example, seasonal changes, rainfall events impacting the system). For example, in an experiment to evaluate the long-term impact of burning on grassland insect diversity, burned and unburned plots are monitored over time, such that the unburned plots serve not only as a burn control, but also as a control for separating the effects of burning from seasonal changes in the insect community.

3.4 Pilot studies There is no substitute for really good planning, including clear formulation of the research/conservation question, and choosing appropriate statistics. Nevertheless, this is still only a first phase—the armchair phase—which should be developed into a concept note of at least a couple of pages, detailing all you intend to do. This concept note can then be used to bounce ideas off fellow researchers, site managers, and supervisors. Then, before embarking on a fullscale sampling programme based on this concept note, you need now to go out into the field and do some preliminary sampling, such as walking a transect, setting a few pitfall traps, collecting some focal insects, etc. Such a feasibility study or pilot study is invaluable, and enables you to go back to the concept note, refine and develop it into a research proposal, and then implement it, knowing, with confidence, that what you have planned to do is achievable in the time allotted to the project. Pilot studies serve several useful functions. Once you have made preliminary decisions on the first ten questions in Table 3.1, it is usually wise to then conduct a pilot study as a preliminary attempt at implementing the study you have planned, in which the first set of data are collected relevant to the objective. Pilot studies are usually of short duration, involve few samples, and may be conducted over a smaller extent than planned for the final study. Pilot studies are used to: • Test equipment and sample methods • Assess if the sampling can logistically be completed in the time available

Coping with the unexpected | 81

• • • •

Assess if sampling can logistically be conducted at the sites selected Practise the implementation of sampling techniques and methods Generate estimates of the variance for conducting power analysis Generate data for planning which statistics it will be necessary/most appropriate to use • Identify unforeseen problems. After the pilot study is complete, the original design may be modified to incorporate the lessons learnt. Pilot studies are almost always extremely useful, and most often indispensable to the success of a study. If the pilot study runs very smoothly, and no alterations to the initial design are made based on the outcome of the pilot study, then the data collected during the pilot phase may also in some instances be incorporated into the final analysis.

3.5 Coping with the unexpected What happens when things go wrong? As ecologists, we should be used to fire, flood, drought, or other natural catastrophes interfering with our carefully planned sampling protocol, or at least expect it at some point in our career. The contents of pitfalls may be raided by hungry hornbills or vervet monkeys, intercept traps trampled by elephants, site markers eaten by termites, and Malaise traps disappear and turn up as kitchen curtains. What do you do if your study is delayed or biased, or you lose samples or replicates as a result of such events? Good pilot studies should help to eliminate many potentially nasty surprises from factors that it is possible to control or manage. Also, try to ensure in advance that the chosen sites will be secure from interference or planned disturbances for the duration of the study. In some conservation scenarios, site developments may be placed ‘on hold’, perhaps via a formal moratorium, pending environmental assessment. However, sadly, not all such events can be anticipated and the worse case scenario may involve a year’s delay for the project, a start from scratch, or a move to a different study area. Such is the life of an ecological entomologist. However, always be on the look out for the silver lining, where disasters may be turned into valuable opportunities to improve understanding or management potential. For example, an unexpected fire across your study area halfway through sampling could turn a well-designed study into not only a comparison of habitats, but also a comparison of insect diversity before and after fire. This raises the problem of missing samples. How many missing samples are acceptable? What does one do with missing data during analysis? Unfortunately, there is no single answer to these questions, and it depends on the sample design,

82 | Designing sampling protocols for insect conservation

and initial sample size, or number of replicates per treatment. Generally one or two missing data points per treatment will not severely affect the analysis or outcome, and the larger the number of replicates to begin with the better off you will be. In cases where, for example, one pitfall is lost from a set of 10 pitfalls that together form the grain of the study (i.e. all 10 pitfalls are used as a single sample), the entire set may have to be excluded from analysis. A set of 9 pitfalls is not comparable with one of 10, because the sample effort differs between them. However, instead of discarding the entire set you could use the average catch across available pitfalls (comparatively insensitive to small differences in pitfall number per set) in a set rather than the total catch (very sensitive to any differences in pitfall number between sets).

3.6 Data management As soon as the objectives of a study start to be defined, the job of data management begins. Data management includes: • Keeping track of which decisions were made and why during the design and planning phase • Keeping record of data collection and data entry dates and error checking • Collating records of observational comments and field notes made while the study was being conducted • Reminders and updates of electronic and hard copy backups of data books and electronic data files; your institution may have formal requirements in place for this important data storage • Keeping track of sample sorting and specimen identification, sample container and specimen labels (e.g. meaning of acronyms or nicknames used and housing of voucher collections and unsorted samples) • Keeping track of exploratory data analysis and statistical analysis data files and results • Keeping track of versions of the manuscripts produced • Maintaining a metadata file that describes the date of creation, location, contents, updates and backup information of all of the above, i.e. of electronic and hard copy data files, samples, and voucher collections.

3.7 Summary To ensure that the conclusions drawn from your research study are as reliable as possible, it is extremely important to plan the study well (Table 3.1). This

Summary | 83

includes a clear, detailed outline of the study objectives (including hypotheses to be tested) and formally designing a sampling protocol. During the design of the sampling protocol, decisions are made about, for example, sample extent, grain and number (Table 3.1). Samples must be well replicated and positioned, so that they are independent, to avoid pseudoreplication. This may involve measuring the spatial autocorrelation of variables of interest across the study area. In manipulative, and also sometimes mensurate, studies, the concept of having controls is important to ensure correct interpretation of treatment or disturbance effects. Pilot studies are used, amongst others, to iron out problems of sampling protocol before the main study is initiated. A well planned study not only ensures that the outcome provides valuable, accurate and novel information, but is reasonably robust to unexpected events while it is being conducted. Planning includes the development of a data management system that will be used throughout the duration of the research, and in many cases for posterity.

This page intentionally left blank

4 Collecting and recording insects

4.1 Introduction: What do you really need to collect? Specimen collection (or recording) starts once the ecological or conservation question has been identified and once the study objectives have been set. The records or specimens collected will constitute your samples or voucher specimens from which to answer the questions posed at the start. The aim is always to collect insect specimens ethically, parsimoniously, and efficiently. In other words, time and effort is directed specifically to answering the question posed. This does not necessarily mean that you are so locked in that subsequent new ideas are precluded. However, any new set of records or samples must be planned from scratch, or should be directly comparable with results already gathered. This point re-emphasizes the importance of advance planning, as well as preliminary sampling, as part of a pilot study (see Chapter 3.4). Any single technique of collecting insects provides only one view on the world. An analogy is a set of windows around a building. Looking through one window (i.e. use one method in one way) provides one view, and when you look through a different window (use another method), you are likely to see a different picture of the outside world. Two adjacent views (two similar techniques) may give you similar but overlapping results, whereas a window on the opposite side of the building (two completely different techniques) will give you a totally different view (very different types of results). The outcome of this is that it is often worthwhile to use complementary sampling techniques. For example, when sampling moths in a wood, the noctuid and arctiid moths may be best sampled with a light trap, while the geometrid moths may be best sampled with a light tower (see below). Where the aim is to sample all the moth species in the area, it will be necessary to use additional techniques, such as treacling and sleeving (see below), as well as searches for larvae and pupae. This returns us to the point that you must be very clear on the aim of the project and on the amount of

86 | Collecting and recording insects

time and financial resources available. Should you focus on a restricted spectrum of insects, ranging from a single order to a single species, your methods can be tailored specifically for this. It is always important to routinely obtain as much background biological information as you can to streamline your approach and efforts. For example, if your insects are in the accessible adult stage during a particular part of the year only, there is no point in seeking them in other seasons. For example, much of the extensive published literature on the seasonality and biology of pest insects and sampling them in crops contains information invaluable for other contexts of sampling insects. Another consideration is that different insect collecting techniques also operate or attract over different distances, which may also be different for different species. Pitfall trapping (see below), sweep netting, and hand-held suction sampling are basically point sampling, whereas Malaise traps will capture individuals over at least several metres. In turn, light traps and pheromone traps have an area of operation that can be enormous (attracting insects from up to several kilometres away). So, if you are comparing, say, different clumps of trees that are within sight of each other, light traps would be inappropriate. Different collection techniques will be variably effective against different species, sometimes even for those in the same genus. Also, any one technique may be effective at certain times of the day or night (different moth species, for example, visit light traps at different times during the night, reflecting their different patterns of activity). Any particular species may be differentially sampled by any one technique according to the age of the component individuals. Most mature adult male dragonflies, for example, can be sampled along the river’s edge, but the young adults of some species are usually not sampled because they move away from the water (even up to 20 km) to mature. There are also differences between the sexes in their response to a particular collection technique. An extreme case is among some moths, where the males are strong fliers while the females are flightless. Finally, bear in mind that for holometabolous insects (i.e. those where the young stages are morphologically and functionally very different from the adult) one is dealing with two functionally different organisms: a caterpillar is very different from a butterfly in both its habitat and its resource use. What this emphasizes is that the different sexes or life stages may require very different collection techniques. So it is important to bear in mind that when one collection technique is used, besides not necessarily capturing different species equally, it may also not capture different sexes, life stages and even, sometimes, different individuals equally. An assessment using a wide variety of techniques, and illustrating the complementarity of the different methods, is given by Clark and Samways (1997).

Named and nameless approaches to data portrayal | 87

At the outset the study must be planned with a clear understanding of the spatial and temporal scale, and then enough sampling conducted to provide statistically sound results (see Chapter 3). Similarly, the collection technique to be used will hone that view. In other words, it is a question of how the sampling is done as well as how much, and deciding upon the equipment needed. While you do not want to take too few samples, you also do not want to take too many. Doing so may involve unnecessary and thus unethical collecting, to the extent that the samples cannot be sorted and analysed in the time available. Obvious as this may seem, it is still surprising to see how often research projects cause stress and even fail to materialize because of an undisciplined sampling protocol (see Chapter 3.3). We now provide a broad overview of the collection techniques available. Inevitably in an introductory text such as this, not all techniques for every taxon and every concept can be accommodated. It is therefore essential to read around your specific topic, carefully scrutinizing recent research papers, to ensure that you are aware of the appropriate sampling options open to you at the outset of the study. This chapter should be read in conjunction with Chapters 2 and 3.

4.2 Named and nameless approaches to data portrayal It is very useful to bear in mind when you start sampling that you are collecting data that can be analysed using two different broad approaches, which from a casual perspective may be described as named vs nameless approaches. To illustrate this point let us say that you sample a particular number of individuals of a moth species, which we shall call (fictitiously) Mothus oneus, in a particular place (where your trap is). At the same time, you also sample another species, M. twous, in the same spot, and a further species M. threeus, as well as other species to which you give names. You have thus collected three and more named entities in one place at one time (Table 4.1). In conceptual contrast, when you ‘number crunch’, plot and analyse these results simply as numbered entities (that may nevertheless have names), they effectively become nameless (as in Figure 4.1). In other words, the coordinates on a graph and the associated statistics are effectively faceless species, and without superscripts, there is no telling which point relates to which species (Figure 4.1). This production of graphs with faceless species has become a problem in recent years with journal space being so limited, and no space available (unless electronically) to list which species are included on any one graph. There are some wonderful numerical and graphical distillations of the data that provide crisp, deductive illustrations for the description of broad ecological or distributional patterns. Yet nameless approaches are

88 | Collecting and recording insects Table 4.1 Named species and abundances of moths sampled using a particular trapping technique, e.g. a light trap. The ‘scientific names’ used here are of course fictitious, and are given simply for illustrative purposes Species Mothus oneus M. twous M. threeus M. fourus M. fiveus Anothermothus primus A. secundus A. tertius A. quartus A. quintus A. sextus Trimothus alpha T. beta T. gamma T. delta T. epsilon Fourmothus unus F. duus

Abundance 21 7 36 231 333 924 428 989 72 4 104 35 098 667 3 268 27 528 7 402 7 002 73 808

not always helpful, or sufficient, for on-the-ground conservation where particular species are the focus. The named species and associated quantitative data can be immensely useful, particularly for other researchers. These associated data, as with the voucher specimens, require careful and accessible storage (see Chapter 2). This can be done in the form of appendices in reports or theses, or as supplementary electronic data, and can be referenced in the scientific paper. This named vs nameless conceptualization is a special but important issue with insect and other biodiversity studies involving large numbers of species. The above points need to be borne in mind when you choose collection techniques, because as with bycatch issues (see Chapter 2.1), you need to be mindful of other opportunities that may present themselves in the future. This can be very important, for example, when comparing biodiversity in an area from one time (one study) to the next (study). In summary, when planning the study, you need to think on the one hand how you will ‘melt down’ your data for the inductive, faceless statistics and graphics, and on the other hand, be aware of how and where you will include the named, semi-raw information. More

Named and nameless approaches to data portrayal | 89 7

Abundance (log)

6 5 4 3 2 1 0 1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 Rank

Fig. 4.1 An example of ‘nameless’ analysis (in this case in the form of a rank abundance distribution used for identifying and describing broad patterns in biodiversity) of the ‘named’ species data given in Table 4.

details on the inductive approach are given in Chapters 3 and 9, but the subject is raised again here to emphasize that you must plan and visualize what you are going to do with the specimens and data right at the start of the project (the software Biota is a very useful tool for biodiversity data management; see p. 373 for details). Because substantial effort (time, cost, and person-hours) goes into collecting insects, wherever possible the maximum information should be extracted from the specimens obtained. We now describe some of the techniques and apparatus used to collect and sample insects. There are many variations on the details of these methods, and many types of apparatus are available commercially and can be found on the Internet. Should you wish to construct some of them yourself, you will find more details of measurements, at least of the more traditional items, in Martin (1977) and Uys and Urban (2006). But always bear in mind that any piece of apparatus may need to be adapted in one way or another for answering your particular research question. Another point to bear in mind is that insect ecology and conservation need not necessarily focus on specific taxa, but on, for example, the particular size of the component species (as large species tend to go locally extinct before smaller species in the same taxon), or on particular functional groups or guilds. Basset et al. (2004), using three complementary trapping techniques (Malaise, pitfall, and yellow pan traps) illustrated this by showing that indicator taxa (see McGeoch 2007) of forest disturbance belonging to selected chewer, parasitoid, and predator guilds had more discriminatory value than other guilds or the use of higher taxonomic levels (e.g. ordinal level). A brief overview of some of the techniques available, and the context of their application are given in Table 4.2.

90 | Collecting and recording insects Table 4.2 Summary of some approaches and methods used to collect or sample insects in ecological studies. See text for additional details Target habitat Some methods available

Comments

Forest canopy Direct access, via walkways, Allows direct, close-range sweeping and foliage cranes, platforms, ropes collection, but often expensive; safety issues are critical Fogging/misting using Accessed from ground or by hoisting machinery insecticides to canopy; falling insects collected on trays/ funnels on or near ground. Use only in calm, dry conditions; wide range of insects sampled Direct foliage samples Obtains insects from twigs or branches by beating or using an aspirator; direct sampling of galls and leaf mines; collection of insects emerging from plant parts using sleeves; mostly targets sedentary taxa Low canopy, Sweep netting Uses standard, coarse nets among foliage to dirshrubs ectly capture free-living insects; fairly specific to certain taxa and life forms; done in calm, dry weather Beating Uses standard horizontal canvas tray, with foliage jarred to dislodge insects; insects individually collected from tray using soft forceps or aspirator; done in calm, dry weather; easy replication Selective individual For large, individually sought insects; collect by collection netting, forceps, insect tongs, with aspirator or with sticky rod; causes minimal disturbance Low vegetation Sweep netting As above; collects wide range of insects that are dislodged relatively easily; can be standardized by certain lengths of sweeps in grasslands/crops Suction sampling Uses suction devices, but must avoid clogging of machine; use only in dry conditions; both widenecked, and narrow-necked (90% ethanol). The glycerol reduces evaporation. Ethylene glycol (antifreeze) is another popular preservative for pitfall traps, although it is toxic to humans, and of course, is not readily available in tropical countries. Make sure that the preservative used does not repel the insect group of interest. The traps are dug into the ground with minimal disturbance to the substrate and the rim left flush with the ground surface. It is generally a good idea to leave the traps closed for a few days before setting, to compensate for the digging-in (or disturbance) effect. Pitfall traps are normally left open for a certain amount of time, usually 3–5 days, although Borgelt and New (2006) provide strong evidence that 7 days is the minimum suitable period for some applications, but it depends also on weather conditions. Pitfall traps deployed in the rainy season may well fill up with water within a day or two, and thus require much more regular emptying, or providing each trap with a roof or shelter as noted below. At the end of this trapping period, the traps are emptied into collection bottles and labelled with pencil on paper inside the bottle. It is generally better to rest the traps by closing them up between the sampling periods, to minimize disturbance effects and to provide time for the researcher to sort, or at least properly store, the captured specimens (sometimes several hundreds or more). After the rest period, trapping can be resumed. This may be done throughout the year if appropriate, or at set intervals: for example, four 7-day trapping periods at four discrete times of the year, representing spring, summer, autumn, and winter. The choice of trapping frequency is dependent on the study objective (Chapter 3). At high latitudes it may of course not be appropriate to sample during the winter. When traps are closed between sampling periods it is essential to mark where they are, because they become overgrown and difficult to relocate. This can be done directly, by marker stakes for example, and/or using fine GPS coordinates. There has been considerable debate as to which type of pitfall trap is best, although choice of trap depends in part on the types of focal insects (or other arthropods). The Majer-type pitfall trap (Figure 4.26) is excellent for ants (although other types are also effective for ants; Agosti et al. 2000). It is essentially a boiler tube, about 18 mm in diameter. It is inserted into a sleeve (made from irrigation pipe or similar) that is sunk permanently in the ground. When the boiler tube is removed, the sleeve is corked up. The sleeve needs to be banged into the ground with a custom-made, very tight-fitting awl. Some researchers working in firm soils have found that the sleeve is unnecessary and the tube can be sunk directly into the ground, but traps may then become very difficult to extract easily. For larger insects such as ground beetles, larger traps need to be used (Figure 4.27). The diameter of the mouth of the pitfall trap has a large effect on the

118 | Collecting and recording insects

Fig. 4.26 Majer-type pitfall trap for sampling surface-foraging ants. This trap is effective for most ground surface-living ants, and excludes large insects and other bycatch. The sleeve consists of a chamfered strong plastic tube (such as irrigation piping) that fits snugly over a purpose-made steel peg. The two are driven into the ground and the peg extracted. The tube is then the housing for a test-tube or boiler tube that is chosen to fit fairly tightly into the sleeve. The trap is then partially filled with a preservative (e.g. 2/3 70% ethanol and 1/3 glycerol) and left operational for a few days, after which the contents are poured out. When the trap is not in use, the sleeve is stoppered. A shelter for the trap, consisting of a roof on four stilts to keep out rain, is an additional option.

Fig. 4.27 Large-diameter pitfall trap that catches a wide variety of insects and other surface-foraging arthropods, both small and large. One plastic cup sits inside another, with the inside one being removed to empty the contents. Various preservatives can be used, with one of the most versatile being 80% ethanol and glycerol (Courtesy James Pryke).

Sampling insects on the ground | 119

diversity and body-size spectrum of insects trapped. Disposable plastic coffee cups have been very popular because they are cheap, easy to obtain, and light to carry. One cup can be sunk as the sleeve and a second one inserted inside the first as the trap. These larger traps are very prone to flooding during rains, and so they may require a shelter, such as a plastic Petri dish on wire legs or a galvanized nail holding the shelter a few cm above the trap. These shelters of course are very vulnerable to mammalian disturbance, including that by humans. Pitfall traps can also be baited. The bait is placed in a separate container, covered with gauze cloth to prevent the attracted insects remaining on the bait, inside the pitfall traps and on a pedestal so that it is above the preservative. Choice of bait depends on the target taxa. Rotting fish or meat can be used to attract carrion beetles, while chafers can be attracted to fermenting fruit. Coprophagous beetles can be attracted to traps baited with mammalian faeces. The quantity of bait per trap should be standardized. As many authors have emphasized, the catch from pitfall traps depends on the activity patterns of the local species, and also on habitat structure, and so the proportionate number of captured individuals may not necessarily reflect the true numbers of individuals from one species to the next. Because habitat structure can have a significant effect on pitfall catches, it is advisable not to use pitfalls to compare habitats with very different structures. But this largely goes for most trapping techniques. Pitfall traps are excellent for point sampling and for comparative sampling in different habitats. However, it is essential that results are made comparable by the traps being open for exactly the same amount of time (e.g. if a 3 day rotation is decided upon, then it must be adhered to). This may require some preliminary sampling to ascertain the best length of time for the trap to be active. Also, all the traps must be flush with the ground so that their trapping efficiency is equal. Trap layout depends on the aims of the project, although our advice is that individual traps be replicated in one spot to improve efficiency (see also discussion in Chapter 3). One way to do this is to place them in a square: say 1 m² for the Majer trap and 2 m² for cup traps. These would not, however, be replicates in a statistical sense, but would be considered as one ‘trap set’. Each set would then be a replicate for analysis. The number of traps in a set will vary, but some workers suggest 10–15 traps is adequate. Some authors have combined pitfall traps with barrier traps, to deflect the wandering insects into the pitfall trap. There are almost as many designs as there are studies using them, with a generalized design given in Figure 4.28. The barrier or fence can be a wooden plank or plastic corrugation, about 2 m long, at each end of which is placed a pitfall trap. These traps are particularly effective for large, fast-walking insects such ground beetles and tiger beetles. As they scuttle

120 | Collecting and recording insects

Fig. 4.28 Polythene barrier trap, 50 m long (above), at either end of which is a pitfall trap (below), being used here to catch large savanna arthropods (Courtesy James Pryke).

across the surface of the ground, they bump into the barrier and then follow it along, only to fall into one of the pitfall traps at either end. Cockroaches and other insects that are shy of light can be captured by placing dark-coloured cardboard or wooden condos into which they retreat as night time ends. These condos should be only about 1 cm high and have an entrance on

Sampling insects in the leaf litter and soil | 121

either end that is funnel-shaped to encourage them in. Traps can also be baited, either with a carbohydrate such as wheatgerm or with something like meatloaf, depending on the target taxa. Traps like this are very effective for insects living on the floor of caves. Finally, in the case of large insects such as grasshoppers in savanna, or tiger beetles, simply the use of a good pair of close-focus binoculars can provide an enormous amount of good quantitative data. Transects, usually 50 m or 100 m long and 4 m or 5 m wide, are walked slowly and all target taxa seen (and verified with the binoculars) recorded on a data sheet. A voucher collection (see Chapter 2.5.3) is maintained, against which you can compare the species you see in the field. Any new species is captured and added to the voucher collection. As it is usually difficult to remember a long list of scientific names, the observed species can be given vivid nicknames that are descriptive in the field. These nicknames are later cross-referenced with the voucher collection to provide the scientific names. This technique only works for taxa (species or genera) that are readily told apart on sight. Even then, capture of individual voucher specimens may be wise, to guard against any mistakes in identification. Watts et al. (2008) developed a method for tracking (without having to collect) rare, elusive, and nocturnal giant weta in New Zealand using tracking tunnels. The tunnels are baited (with peanut butter in the case of weta) to attract the insects inside them, but as they enter the tunnel they are obliged to walk through an ink ‘footbath’. As they progress to the bait in the centre of the trap, they leave inked footprints behind on the white card base (Watts et al. 2008). The method has been so effective for weta that it is worth trying out on other large, ground-living insects.

4.8 Sampling insects in the leaf litter and soil Sampling insects below the surface of the litter or soil generally goes hand in hand with sampling other arthropods and even other invertebrates (New 1998). Together, they all constitute the litter or soil fauna. Insects living in logs (saproxylic species) can often be classed with litter insects as they can be similar in taxonomy and sampled in the same way. The most commonly employed technique is that of the Berlese–Tullgren funnel (Figure 4.29). It can be used for litter, soil, log, and bark samples. The principle is that a sample is placed over a coarse gauze sheet (about 5 mm mesh size) and a 60 W light bulb used to provide both light and heat to drive the fauna down into a funnel, at the bottom of which is a collecting bottle with 70% ethanol. It is usually necessary to place a small gauze mat with a smaller mesh size (about 1 mm) in the centre of the coarse gauze to prevent litter and soil samples from falling into the funnel and dirtying the collecting bottle.

122 | Collecting and recording insects

Fig. 4.29 (Upper panel) Bank of Berlese–Tullgren funnels with a series of six 60 W light bulbs above each funnel. (Lower panel) Bank of funnels with the canopy holding the light bulbs removed to show the funnels and the collecting bottles at the base containing 70% ethanol underneath (Courtesy James Pryke).

The samples can be placed in the funnel according to a set mass or volume. This makes quantitative comparisons between sites possible. However, it is essential to bear in mind that the results can be highly biased if the samples are sometimes taken after rain and at another times after a dry spell. In the case of log samples, the woody material can be broken up by hand and then placed inside the funnel. As one usually works with replicated samples the funnels are normally constructed in rows (Figure 4.29). This also means that extraction of the insects and other fauna from several samples can take place at the same time. Working on the same principle as the Berlese–Tullgren funnel, and where electricity is not available, an alternative technique is the Winkler bag. This is simply a gauze-covered frame shaped into a funnel at the bottom and at the end of which is placed a collecting tube with 70% ethanol. Inside the frame is placed

Sampling insects in the leaf litter and soil | 123

a bag made of 4 mm mesh that acts as the retaining sack for the litter sample. The whole of the Winkler bag is suspended from a tree or a frame. As the litter dries out, the insects and other invertebrates move down and move through the mesh of the inner sack and tumble down into the collecting bottle at the bottom. Quantitative data can be obtained by placing a set weight or set volume of litter or soil in the inner sack. The whole bag is lightweight and can be folded up very easily, making it ideal for expeditions to remote areas. A ground emergence trap can be used to sample emerging insects from the litter layer and soil. It is particularly useful at the start of the summer season and after rains in dry areas. The trap consists of a tent made of black gauze pegged to the ground with a collecting bottle placed in the top corner to collect the insects, especially flies, emerging from the ground. A similar principle can be applied to logs using trunk emergence traps. These are constructed by wrapping the entire circumference of a random 30 cm long section of the log (although never closer than 50 cm to the end of the log) and the trunk of snags with polypropylene weed barrier cloth, which permits moisture and air to pass through, but not light. Pieces of strong wire are inserted into holes in the log to keep the cloth away from the log, and the cloth sleeve wired at either end to confine emerging beetles. Foam or carpet underlay is placed between the cloth and the trunk, under the wire seals, to accommodate the rugosity of the bark. The insects are collected in a 250 ml clear plastic bottle attached by a screw thread to a lid at the top end of the sleeve, and half-filled with 50% propylene glycol. Beetles emerging from the log fly towards the light and are caught in the bottle (Johansson et al. 2006). Finally, it is necessary to mention termites as a special case (the same principles also often apply to ants). They are eusocial insects and require special collection techniques (Jones et al. 2005 provides an overview). Of course not all termites live in the soil; some are arboreal or live inside rotting timber. With termites it may not be necessary (or appropriate) to sample individuals. Instead their mounds can be recorded, including mound size, density, and dispersion. Termite mounds can also be dug up, but bear in mind that searching for the queen can involve a very deep dig indeed, as she may be several metres below the soil surface. Jones et al. (2005) recommend that a quick way to make a rough estimate of termite population size is to take a sample of known volume of the mound, extract and count the termites, and multiply up to the volume of the mound. The problem is that mounds can be irregular in shape. This is partly overcome by the mounds being assumed to be regular shapes such as hemispheres or cones (or more accurately using photography and image analysis software). The initial sample must be taken quickly, before the alarmed termites retreat further inside the nest. They can be sorted from the nest material by hand

124 | Collecting and recording insects

searching or by floating the individuals out in water in a flotation tray. Caution is required because termite occupation of the upper part of the mound depends on the time of day, temperature, and whether it is a dry or wet period.

4.9 Sampling aquatic insects Insects are abundant in the aquatic environment, particularly where the water is shallow and with an abundance of water weed. Sampling under these conditions requires a very strong dip net (Figure 4.30). The most versatile water net is triangular, about 30 cm across at the far end, with a frame made of stainless steel. The bag is best made from stainless steel with 1 mm gauge mesh. Obviously the handle must also be very robust. Such a net can be pushed vigorously into water weed to dislodge insects. Many insect larvae and other invertebrates live on the bottom of water bodies. Sampling of these is well summarized by Southwood and Henderson (2000). They point out that there are five principal variables that affect the choice of sampler for bottom fauna: (1) the type of animal being sampled; (2) the nature

Fig. 4.30 Dip net used for catching aquatic arthropods: it must have a sturdy frame and tough gauze (Courtesy Ricky Taylor).

Sampling aquatic insects | 125

of the bottom substrate, whether soft or hard; (3) the speed of the current; (4) the depth of the water; and (5) the objective of the study. From a conservation perspective, rivers and wetlands can be assessed for ecosystem health based on the composition of the benthic macroinvertebrates (Hauer and Lamberti 2007). Each taxon (usually family in the case of insects, but a higher taxon for most of the other invertebrates) is assigned a sensitivity score (see Gerber and Gabriel 2002 for an example) and the total score reflects the condition of the water body. A much more robust and meaningful comparative score is the average score per taxon (ASPT), which is simply the total of the sensitivity scores at any one site divided by the number of taxa recorded. Smith et al. (2007) provide a worked example and application of this approach. To obtain the macroinvertebrate score for river assessment, the dip net mentioned above is used as a kick net, where the net is held upright, and then, immediately upstream of the net, the stream bottom is stirred up by moving your foot around, until all the bottom is stirred up in a standardized set of movements. The net is then searched to remove excessive debris and large insects removed with soft forceps. The contents can also be tipped into a white tray of clear water. Specimens can be sorted in the tray using a fine sable-hair brush and specimens removed with soft forceps and, after being counted, returned to the water, or, where needed for later identification, placed in vials with 80% ethanol. This technique, when employed over a set time period (usually 15 min) or over a particular area of water, provides much quantitative and comparative data. When a set time period is used, all microhabitats at the site being assessed are included in the 15 min period. As species-level assessments provide much more sensitivity than the use of higher-taxon surrogates, and for measures of ecological integrity, selected individuals of insect larvae can be reared through to adulthood in specially adapted aquaria. This method enables matching of any particular larva to an adult, and is particularly useful in the case of dragonflies, details of which are given by Rice (2008) and Samways (2008). Emerging aquatic insects can be sampled, even quantitatively, using an aquatic emergence trap placed over the surface of the water (Freitag 2004). As with many terrestrial traps, such a trap is sampling continuously once it is in place, and has the advantage of collecting aquatic insects that have emerged from the water as adults and might otherwise have dispersed. It is essentially a fabric tent that is placed over the surface of the water, usually adjacent to the bank. There are various designs depending on intended use (see Freitag 2004), with the simplest one being a frame of plastic tubing and a nylon tent that is floated at the shore to sample emerging dragonflies (Figure 4.31). The advantage of this system is that the larval ‘skins’ (exuviae) are kept on the inside of the tent and can be matched to the adult that is also captured in the tent.

126 | Collecting and recording insects

Fig. 4.31 Aquatic emergence trap, being used here to sample emerging adult insects. As the insects leave the water to become adults, they fly up to the top of the tent where they are directed by a Perspex funnel into a chamber made out of two plastic jars facing each other, with a hole cut into the side of the top one. If the insects need to be kept alive, no preservative is placed in the chamber. Otherwise, a preservative such as 70% ethanol is introduced. (Courtesy John Simaika).

Various other methods are also available for sampling bottom fauna, including removal of set amounts of substrate using, for example, a grab, which is a piece of apparatus which acts as a pincer to literally grab a standardized amount of bottom substrate. Various types of trawls and dredges are also available for sampling along set distances across the bed of still water (Southwood and Henderson 2000). Many of the vast number of techniques along these lines have been developed for specific purposes. Some, such as trawls and drags, can be very disturbing, even destructive, of the habitat, and must therefore be used with caution in a conservation context. Some may only be partially effective, with certain species escaping from the haul. A very useful method for sampling in still and clear water, which is very non-intrusive (and therefore very effective for determining exactly where in the water body and where among plants certain insect species are living), is by diving using underwater scuba gear. Sampling can be done in various ways, with the lead originally coming from coral reef assessments. Sampling can be along set transects under the water (e.g. a 50 m tape measure can be laid out and individuals at each 10 cm can be recorded; or, individuals can be collected using the insect tongs mentioned in section 4.4 and individual positions recorded). This method can generate much detailed and accurate information relatively quickly. The deployment of scuba gear was used, for example, to assess where trout food

Sound recording | 127

was distributed around a reservoir—as it turned out, this was in the 0.5 m weedy margins (Samways et al. 1996), leading to trout fishermen immediately changing where they cast their flies!

4.10 Sound recording Many insects produce sound. This can be very useful for various conservation assessments, especially as many orthopteran species produce high-intensity sounds, and, as a group, they tend to be sensitive to landscape change (see Samways 1997a for a review). Recognition of the specific nature of orthopteran song and the development of lightweight, tight-focus recording techniques has led to incorporation of sound recordings of these insects into taxonomic analyses and evaluations. Flowing from this, the use of insect song can be developed into assessments of habitat quality. Table 4.3 gives an initial categorization of song types. The relative proportions of these, for example, can give an indication of how the landscape has changed in favour or not of the various species. Methods are currently being developed using digital spectrograms to bring much more detailed focus to these acoustic assessment methods. When recording insects, it is important to bear in mind that their song, with its high-frequency components, rapidly attenuates in vegetation, especially the broad-leaf variety. This means that one of the golden rules of recording insect song is to make sure that the microphone is as close to the singing insect as possible. There is a trade-off here, because the closer you get the more likely Table 4.3 Song characteristics for major taxonomic groups of Orthoptera (from Riede 1998) Taxon

Spectrum

Frequency range (kHz)

Infrastructure (human ear terms)

Tettigonioidea

Wide Narrow Narrow

5–101 1–101 1–11

Any Note and chirp Trill

Wide Wide Wide Narrow Wide Wide

2–40 2–40 2–40 3–5 2–40 1–8

Zip, buzz, and rattle Tick and rattle Tick and rattle Trill Rattle Rattle and trill

Grylloidea Acridoidea Gomphocerinae Oedipodinae Acridinae Hyalopterix Romaleinae Pneumoridae

128 | Collecting and recording insects

the insect is to be disturbed and stop singing. One way to overcome this is to suspend the microphone from the tip of a fishing rod and lower it over the singing insect. The insect is far less disturbed when a small object descends from above than when a bulk approaches from the side. Be careful not to break or crunch any vegetation underfoot, as this gives out a sharp, transient sound to which orthopterans are very sensitive. For nocturnal singing insects, and after recording their song, you may wish to retain the specimen, in which case you can search for the insect in the vegetation with a head torch and catch it with the insect tongs mentioned in section 4.4. Use of a parabola, and especially the immensely useful bat detector, for capturing insect songs is generally acceptable for most ecological and conservation applications, especially as most of the insect’s energy is put into producing sound around the carrier frequency, which includes the audible range and, with many bush crickets (katydids), also or even exclusively in the ultrasonic range (Hung and Prestwich 2004). Diwakar et al. (2007) have even shown that an experienced listener can accurately record the various songs of audible crickets for conservation assessment purposes. Simple chirp counts may be used at low densities of grasshoppers, but may cause confusion when large numbers are present, because of difficulties in distinguishing the songs of individuals or species. However, recent developments in automatic recognition bioacoustic techniques, by which parameters such as chirp length and pulse length can be extracted from recordings (Schwenker et al. 2003), have led to improved possibilities for species identification and enumeration. Work on British grasshoppers (Chesmore 2004) showed that species identifications can approach 100% accuracy under low noise conditions, using identifications based on artificial neural networks. Gardiner et al. (2005) considered that acoustic techniques have been under-exploited for monitoring Orthoptera, and that these have important roles in identifying the presence of (1) cryptic species, (2) non-macropterous individuals (which may not be amenable to flushing), (3) species that are very rare and otherwise likely to be overlooked, and also in habitats where more traditional sampling methods are difficult to implement. Advances continue to be made rapidly, with moves toward automated identification systems incorporating abundance estimates. Riede (1998) and Riede et al. (2006) have given much relevant background to the subject, emphasizing the values of acoustic parameters in inventory, as a firm estimate of species richness that helps to overcome gaps in formal descriptive taxonomy, and as tools in monitoring habitat use (and therefore habitat quality) by Orthoptera.

Summary | 129

4.11 Summary It is important to bear in mind that any one technique will only collect a particular subset of fauna. Thus, depending on the research question being posed and the specific aim of the project, it is often wise to use a variety of techniques for different suites of taxa. For comparable catches and therefore comparable data, techniques must be standardized, e.g. for placement, design, duration, bait type and quantity used. Individual traps or technique applications should generally not interfere with each other (e.g. be placed too close together or repeated to soon after each other). Care must be taken to construct, install and apply each trap or piece of equipment so that there is minimal sampling error and disturbance to the environment. Care should be taken when interpreting the meaning of number of individuals of each species caught, for example, as either abundance or density. In most cases trap-based estimates are density estimates (number of insects per unit area or volume), rather than actual abundance estimates, because most trapping techniques have a fixed area over which they operate. Time of day, season, and weather conditions generally have a significant impact on catch, as does habitat structure and landscape topography, and these must be standardized. More than a single technique may be necessary to optimize inventories (called sampling sets). Pilot studies are important for testing the efficacy of a technique, unless it is a standard, well-known, tried and tested method. And lastly, do bear in mind that replication must be done in a way to avoid pseudoreplication, as mentioned in Chapter 3.3.4. As a corollary, we should mention that there are a multitude of techniques, variations, and minor modifications of specific techniques for collecting insects. Most of the traditional ones are discussed by Southwood and Henderson (2000). However, such techniques are often modified by individual researchers and for individual studies, to suit the specific purpose and context of that study. In this chapter we have provided only a broad overview of a range of techniques available and commonly used examples of each.

This page intentionally left blank

5 Measuring environmental variables

5.1 Introduction: selection of variables Insects respond to a range of conditions around them. Many insects have precise requirements for food (such as a particular host plant or dung type) and less readily quantified environmental requirements, such as suitable microclimate. Their incidence, abundance and distributions are governed strongly by such features. These conditions (broadly, environmental variables, EVs) may be determined by other organisms (biotic variables, such as interactions with plants and natural enemies), the physical world (abiotic variables, such as temperature or stream flow), or a combination of both (such as litter depth). Abiotic and biotic variables may combine to strongly influence the seasonal activity patterns of insects, and favourable conditions dictate the rate and timing of such development. Less predictable events—such as fires or floods—may be superimposed upon these patterns and provide opportunities for insects to exploit novel conditions and expand their activities or, conversely, cause direct mortality. The importance of timing of weather events is clearly seen, for example, in the desert, when flowers appear in profusion only some time after the rain, making conditions appropriate for pollinators that then appear rapidly and in unexpectedly high numbers. The focus of this chapter is on measuring EVs that affect insect life (including activity patterns, survival, natality, population dynamics, and interactions with other species), both to its benefit and to its detriment. It is the balance of these two that is often critical for conservation, where of course we wish to create and sustain conditions that are advantageous to the species in question. Indeed, a starting point in conservation is that physical and biological conditions must be optimal for survival and success of the species. In turn, for long-term conservation, environmental conditions must be managed so that they remain optimal. This may involve interventions such as deliberate burning, selective grazing by mammalian herbivores, or the removal of alien plants. A note of caution is

132 | Measuring environmental variables

perhaps needed, because many insects of conservation interest are ecological specialists, and as a consequence are rare and potentially vulnerable. Many species are known from few, even single, sites that are presumed to be remnants of a previously more widespread range. Because we have no other model, we assume that the environmental conditions of the occupied site are optimal for the focal insect. Management then seeks to define and replicate these conditions to provide optimal habitat for the species. However, the insect may be only just surviving in the last remaining but impoverished habitat, rather than thriving (Davies et al. 2006). Subtle appreciation is needed to understand this, as otherwise we run the risk of replicating suboptimal habitat conditions for species. As stated by Wikelski and Cooke (2006), successful species conservation relies on an understanding of the physiological responses of species to their changing environment. Achieving such understanding will involve both measuring the environment and understanding how environmental change affects the physiology of the species. The effect of EVs on the physiology of the insect (other than direct mortality) will then translate into changes in, for example, individual behaviour, fecundity, and generation time, that will in turn translate into changes in population size, distribution, and species assemblage structure. Understanding how insects respond to their abiotic environments has become particularly important in the face of the current rapid rate of climate change (Chown and Nicolson 2004). The role of EVs can be illustrated using the reintroduction of insects into a habitat after the local population has become extinct. This procedure is predicated on the causes of earlier decline being known and countered so that the loss will not recur. Elsewhere the species may be surviving, and hopefully thriving, and it is there that we can assume that conditions are at least suitable, if not optimal. These suitable conditions are provided by a combination of site parameters, such as adequate patch (habitat) size, good patch quality, and minimal patch isolation. This would be a source habitat, where the species is breeding and from which individuals are able to disperse to establish in adjacent patches. This contrasts with a sink habitat, where conditions are suboptimal and the local population is as a consequence less viable, with few to no individuals being produced and surviving in the patch. The existence of source habitats is especially important in the long term, because if conditions in at least some patches are not optimal, the species may well go extinct. Site features, such as those above, are one category of EV. Resource supply is another category, and ‘habitat quality’ may be regarded in terms of the coincidence of the critical resources and a climate regime suitable for the growth, maintenance, and reproduction of the insect species.

Introduction: selection of variables | 133

The identification of source habitats is an important first step to understanding and being able to measure what are the suitable or optimal environmental conditions for a species. It is also here that as many as possible of the important EVs should be recorded. These data then provide a yardstick against which other habitats (suitable or unsuitable for the focal species) can be compared. It will most likely be necessary to measure these conditions over a whole year, and/or at critical times, such as at high water levels or during droughts in marshland. When comparing conditions between one patch and another (e.g. a source habitat and a sink habitat), or trying to understand what drives seasonal and longer-term population dynamics of a species, the key question becomes, which EVs should you measure? In turn, if the goal is to maintain habitat quality in good condition over the long-term, the question becomes, which, when, and how often, should critical EVs be measured? The measurement of EVs is also common in assemblage and community studies, where the question of interest has to do with which EVs are responsible for patterns of species richness, abundance, or community structure. The choice of which EVs to measure, when, and at what interval, will depend on the precise ecological question you are asking or the conservation approach being taken. It is all too easy to attempt to measure as many EVs as possible, without understanding the rationale for doing so. Species are likely to vary in their requirements, and more than a single, often interacting, EV is usually important to them. Indeed, EVs often interact with each other in a non-linear fashion. For example, temperature and relative humidity in a microhabitat may increase together until a point is reached where the grassland habitat dries out, at which time temperature may still continue to increase but humidity drops. Different species respond to these changing conditions in different ways. In the case of drying grass, one species of bush cricket, for example, may move to neighbouring bushes while another may move to longer grass. This is precisely why it is so much more valuable to use species rather than higher taxonomic levels when monitoring environmental conditions. A range of species will display a range of sensitivities to the various and changing conditions surrounding them. Where your study involves whole assemblages or communities, or even particular food webs, you will need to ascertain which EVs shape such groupings, i.e. which EVs are associated with high and low species richness, different assemblage structures, abundances, and interactions. Elevation, latitude, and longitude are often measured as relational (independent) variables, especially in studies conducted over broad spatial scales. However, these are not true EVs. Rather they are used as surrogates (or proxies) for a suite of abiotic EVs that are well known to vary in fairly predictable ways

134 | Measuring environmental variables

across gradients in elevation, latitude, or longitude. Similarly, slope and aspect (which can be measured with a clinometer and compass respectively) may be surrogates for drainage conditions, wind exposure, timing of exposure to direct sunlight, and amount of sunshine received daily. Wherever possible it is better to measure EVs directly than to rely on surrogate measures. For example, elevation is a generally good surrogate for temperature. However, fine-scale variations in temperature (microclimates) across an elevational gradient are often not well predicted by a measure of evelation. Deciding on which EVs to measure, and when, is the subject of this chapter. More broadly, you may need to choose those EVs that characterize, for example, ‘high-quality indigenous grassland’ or ‘high-quality ponds’, rather than necessarily considering the representative focal species living there, to define a broad suite of umbrella values that must be sustained. Much of need for selection of EVs will initially be site-focused, and the nature and characteristics of any particular site may dictate the features of interest, e.g. closeness and size of rocks on a hill in a savanna landscape. It is often worthwhile taking a series of digital photographs of the features of likely interest (not least to act as a memory aid in the final selection of which particular features to investigate) as a record of the site, and as a foundation for assessing changes in the future.

5.2 Conceptualizing environmental variable data Before discussing the ‘what’, ‘how’, and ‘when’ of measuring environmental data, let us plan ahead and visualize where you are going to enter these data in the data matrix relative to the insect data itself. The important point here is that the insect abundance or diversity data (see Chapter 9) are often collected at the same time and, of course, in the same place as the various EVs. EVs can therefore be considered relational data because they relate to the insect abundance or diversity data, and usually during analysis one relates the insect data to the EV data in the hope of finding a relationship or correlation between them. Ludwig and Reynolds (1988) provide an excellent and highly recommended introduction based on a data matrix view (see also Chapter 9.3 on the assemblage matrix and relational data). In principle, the analysis of insect data is based on measurements of abundance of species across sampling units (SUs) (see Chapter 3.3). A SU is a discrete, sampled part of a habitat. It may be the number of insect individuals in a particular quadrat, transect, unit of soil, volume of log, volume of water, a particular number of fruits, or so many leaves, etc. The SU is your basic working unit in the matrix. This matrix is arranged with SUs listed across

Choosing the environmental variables | 135

(columns) and species listed down (rows) (a simplified example is given in Table 5.1; see also Chapter 9). The important point here is that at the EV matrix is constructed with the same columns as the species/assemblage matrix (for convenience, usually at the bottom of the data matrix; see Ludwig and Reynolds 1988) so that analyses can be conducted by making comparisons of rows and columns across the two matrices (Table 5.1). You may also wish to have a seasonal perspective, with samples from spring, summer, autumn, and sometimes winter, when influential EVs also vary considerably. This is strongly advised as a minimum basis for understanding the life histories of such short-lived animals. In this case, prepare four data matrices, one for each sampling period. If time allows (because remember that you have to sort all the samples and enter the data into the matrix), you may wish to sample twice during each of these quarterly periods. In practice, a good compromise is to sample six times a year, i.e. every two months. Whatever time period you choose to make your species assessments, the same must be done for the EVs. Importantly, for sound statistical analysis, every SU should as far as possible have its own independent measure of each EV. Depending on the ecological or conservation question, it may be appropriate to focus on certain times of the year: for example, sampling intensively in summer and perhaps not at all in the winter. Because of the fine-grain nature of insect distributions within their overall habitat, tempting surrogates (such as using temperature data from the nearest weather station, or even several metres away) can sometimes be of little value. To achieve broad spatial representation, many one-off samples of species and EVs are often used in preference to repeated sampling at fewer localities (an optimal compromise may in some cases be to sample a few sites several times and the reminder only once). Again, the approach adopted must be driven by the conservation or ecological question that you have decided to focus on (see Chapter 3). And this is where you must be very astute with your environmental data. Do not just measure EVs for the sake of it, but rather measure those that are likely to be meaningful. Of course, when the final analysis of these data is done, some of the EVs will turn out to be important whereas others may not.

5.3 Choosing the environmental variables Deciding which EVs to measure is rarely easy, and the difficulty of choice is rarely mentioned in textbooks. One of the problems is of course that in some cases you do not know which EVs are indeed important until you start measuring, gathering, and analysing the data. In some cases one may be guided by

Table 5.1 A subset of a data matrix in which species and environmental data are entered relative to the sampling unit (SU) in which they occur. In this example, there are 18 SUs. Each of these SUs may, for example, be a quadrat (or the pooled data from say, four quadrats or four pitfall traps). In this matrix there are six SUs (six replicates) in each of the three different habitats(A–C) that are being compared. These habitat types in turn would have to be replicated (to avoid pseudoreplication), say three to six times, for statistical analysis (leading to a larger matrix). In this example only five insect species were sampled, but of course it could have been any number (n). Species 2 is clearly rare (N = 9 individuals) in the sampled area and it was not sampled at all in habitat B. Four environmental variables (EVs) were measured here. EV1 is percentage shade caused by tree canopies over the focal SU at midday. EV2 percentage bare ground, and inversely proportional to this is EV3, percentage ground cover. Also related is EV4, which in this case is maximum height of ground cover. This data matrix is the starting point for all subsequent hypothesis testing and analysis, whether simply descriptive, statistically univariate, bivariate, or multivariate. N = the total number of individuals per species across SUs and habitats Habitat A

Habitat B

Habitat C

N

Sampling Units (SUs) 1 Species

2

3

4

5

6

7

8

9 10 11

12

13 14 15 16 17

18 (. . .n)

1

14

8 21

4 16

9

1

0

0

1

2

0

2

0

0

1

0

0

0

0

0

0

0

0

0

0

2

2

2

1

1

9

3

7 14

7

6

7

2

28 24 32 17 31

27

6

4

4

3

2

7

228

4

2

0

3

4

2

7

0

0

0

0

0

0

70

14 12 12 14 10

18

1435

5 S= number of species

S

Environmental variables (EVs)

2

107 99 78 111 123 166 4

4

1

70 75 80 75 75

2 3 4 … n

4

9

7 11 14

9

105 115 121 97 91 142 4

3

3

4

4

3

75

60 65 60 55 60

58 62 61 57 40

67

28 24 27 27 26

42 38 39 43 60

33 7

7

4

8

4

7

4

8

7

47 40 22 63 71 108

3

4

4

4

4

4

60

80 80 80 85 75

80

26

84 77 81 80 81

82

72 76 73 73 74

74

16 13 19 20 19

18

12 22 19 21 17

13

5

5

4

5

4

6

427

Choosing the environmental variables | 137

literature on related taxa and sometimes even in similar climatic areas or disturbance types. Dung beetle assemblages, for example, are well known to be sensitive to shade and a study on dung beetles may consider canopy cover or light intensity as an EV to be measured. Soil temperature just below the ground surface has been shown, for example, to be an excellent predictor of activity patterns of the invasive Argentine ant (Human et al. 1998). EVs are chosen according to the question you are asking. Say, for example, you are studying the effect of woody invasive alien plants on ground invertebrates. One EV that you would measure is percentage shade cover. In contrast, wind speed at the particular time you are visiting the site may be largely irrelevant to ground-dwelling invertebrates. In contrast, where you aim to measure flight patterns of butterflies, moths, or flies, it becomes vitally important to measure prevailing wind, cloud cover, and temperature conditions. You would therefore need to measure flight activity of the butterflies and at the same time and place measure wind speed and other EVs. Australia’s Golden sun moth Synemon plana, for example, has very finely timed flight activity. Males fly only in calm conditions, in bright sunlight, with temperatures above about 20 °C, without rain, and for a few hours in the middle of the day (Gibson and New 2007). To emphasize this important point, let us take another, rather different, example. Imagine that you have sunk pitfall traps into the ground. Should you measure soil or air temperature, and if so when and how often? Start by thinking it through. What relevance would a point measurement at any one moment in time be to the current pitfall catch? This question applies to both soil and air temperature. Insects and other invertebrates caught by pitfalls may be dependent on the temperatures last night or yesterday afternoon, and in fact perhaps even 2 weeks ago. It may have rained in the night, perhaps stimulating high activity. Present insect activity is generally the net result of the various influences and impacts of both previous weeks and previous seasons. It is critical that you work through which EVs are likely to be important and whether measuring them the way you are is appropriate and will provide a sensible, meaningful answer. This ‘thinking through’ is of course also part of hypothesis formulation (Chapter 3). Furthermore, bear in mind what is critical for one species may not be so for another. A salient reminder of this is the tropical African ground beetle Africobatus harpaloides that tolerates only a 0.9 °C change in daily temperature before dormancy is induced (Greenwood 1987). This illustrates just how critical temperature is for some species. Unfortunately this sort of physiological information is not yet available for the vast majority of insect species (see Chown and Nicolson 2004 for an overview of insect physiological ecology).

138 | Measuring environmental variables

It helps considerably to understand how the insect results relate to EVs after entering your EV data into the data matrix. In Table 5.1, EVs 2 and 3 are the inverse of each other simply because where there is no vegetation cover, there is bare ground. Other EVs may not be so directly equivalent. Air temperature 50 cm above the ground may be related to height of the grass sward, yet the relationship is unlikely to be linear. For example, the temperature will be different, but not necessarily proportional, when the grass is, say, 5, 10, 20 or 50 cm tall. This is because the height of the sward itself may affect the microclimate on the ground, and influence the vitality and activity of epigaeic insects such as certain beetles and ants. Selecting EVs is done a priori. Only when the analyses have later been completed will it become evident which EVs are statistically significant. A pilot study can be very helpful for deciding which EVs to measure in the full study and which can be left out. Rarely is one EV alone the sole driver of insect presence/absence or abundance. Nevertheless, one EV may be a key factor, as may two or three EVs combined. It is important for conservation to determine which of the EVs are significant (Figure 5.1). EVs that are positively and significantly related to insect population size, total abundance, or species richness would be encouraged in management plans, whereas EVs that are negatively related to insect vitality factors would have to be removed or reduced. Identifying a key threat and removing it can lead to an immediate recovery of some insect species.

Fig. 5.1 Before it was rediscovered in 2003 the Ceres streamjack damselfly Metacnemis angusta was feared extinct, having last been seen in 1920. To this day, it is still only known from one pool cleared of invasive alien riparian trees, in a river braid, with the water plant Aponogeton and cleared of invasive alien riparian trees. The point here is that the key threat was identified: invasive alien plants completely shade out the habitat. By addressing the key threat, and restoring appropriate environmental variables, especially sunlight and growth of indigenous vegetation, the species recovered.

Choosing the environmental variables | 139

This was shown categorically when invasive alien woody plants were removed and highly threatened damselflies immediately returned to the site (Samways and Taylor 2004). Sometimes, when you measure specific EVs in the immediate vicinity of a large, mobile insect (such as shade cover, air temperature, and water flow rate close to a perching dragonfly, or when there is a key impact such as road dust on parasitoids; Figure 5.2) you may find that your EVs account for as much as three-quarters of the variance in the insect data. However, for many insect–EV interactions, such high explanatory power (strong correlations between the insect data and EVs) is seldom obtained. Nonetheless, weak correlations may be significant, suggesting that important EVs remain unmeasured. Selecting various EVs or certain combinations of them for statistical testing of significance will often single out which are the really important ones. However, do bear the following three issues in mind: (1) The EVs chosen may not be the only important ones, and other important EVs, both abiotic and biotic, may

Fig. 5.2 The line of citrus trees adjacent to the dust road suffered defoliation (above, arrowed) as a result of a severe Red scale Aonidiella aurantii (bottom left) infestation. Dust is a key environmental variable inhibiting the searching behaviour of the parasitoid Aphytis africanus that normally keeps the Red scale under control.

140 | Measuring environmental variables

not have been measured. (2) The abundance or diversity of your focal population or assemblage may be influenced by, say, a pathogen or competition (biotic interactions), that may, in turn, be responding to different EVs than the EVs to which the focal subject is directly responding. (3) Finally, be very aware of autocorrelation, where different EVs are inevitably spatially or temporally associated. Temperature and shade, for example, usually fluctuate together over both space and time. This means that these two EVs are not independent of each other, and during analysis must be dealt with carefully and usually separately (refer to a general statistics text on the subject of colinearity, or non-independence, among relational variables). Similarly, because different EVs are measured in a range of, often very different, units (e.g. percentage cloud cover, mm of rainfall, degrees Celsius temperature), EVs may need some form of transformation before inclusion in a statistical test (as in fact do insect abundance and richness data) (see Crawley 1993 and Quinn and Keough 2002). A list of possible main and commonly measured EVs to consider including in your study, where relevant, is given in Table 5.2. As a general rule, where possible try to obtain quantitative measures of the EV of interest rather than using a qualitative measure. Sometimes, qualitative estimates, or categories of condition, are the only way to measure the EV of interest. However in many cases quantitative alternatives are available. For example, if given the choice between measuring light intensity in a forest patch vs estimated percentage canopy cover (Figure 5.3), choose light intensity (Figure 5.4) because it will provide a more robust (and often more accurate and sensitive) measure that is easier to deal with statistically when it comes to analysing the data. Similarly, for example, weigh standardized area-based samples of leaf litter to establish litter density, rather than estimating litter cover as, say, ‘high’, ‘medium’, or ‘low’ (the latter provides a categorical EV and the former a continuous one). Basically, wherever possible avoid human subjectivity when measuring EVs. Some general principles for measuring EVs include: • It is better to choose a wide range of EVs, several of which may be complementary. Indeed, measuring a wide range of EVs is often not actually a major practical issue because so many can be measured simultaneously with increasingly sophisticated and affordable electronic instruments. • It is essential to replicate your measurements (replication) as far as possible in the same way (and at the same spatial and temporal sampling points) as the actual insect samples. Several measurements can also be taken in the same microclimate or habitat patch, and the average of these measurements can be used in the analysis. In this case the multiple measures are not

Choosing the environmental variables | 141 Table 5.2 Some essential environmental variables (EVs) to consider measuring when undertaking a study in insect ecology and conservation Category

Comments

Abiotic environmental variables Soil pH

Can be determined with a pH meter

Temperature and moisture

Can be measured with an appropriate probe; relevance to the study must be considered very carefully as the insects and other arthropods present may be present as a result of past temperature and moisture events, e.g. the point of sampling may have gone through a climatic bottleneck in the past such as a high temperatures and drought, or inundation

Compaction

Can be measured with a penetrometer

Physical and chemical composition

Soil samples must be fresh; essential to consult an expert; for introductions to techniques: see Tan (2005) and Carter and Gregorich (2007)

Water pH, dissolved oxygen, and conductivity

Can be measured with a commercially-available water analyser, which must be carefully calibrated; for background information see Hauer and Lamberti (2007)

Turbidity

Traditionally measured with a Secchi disk; water samples can be assessed using a portable turbidimeter;

Pollution

A specialized field; for an introduction to techniques, see Nollet (2007)

Above ground Temperature and humidity

Temperature probes are available, but it may be more meaningful to have a continuous set of data from a data logger or even a local weather station; remember that the current insect assemblage will be strongly determined by past temperatures, especially adverse conditions. Similar issues arise with relative humidity, which can be measured with electronic hygrometers. Insect distributions can be defined by a combination of temperature and humidity; for example, high temperatures and low humidity can be a lethal combination, and measured as the Saturation Deficit Index (see text). Microclimate is generally highly relevant to insects, i.e. the climate within the specific habitats or substrates in which they are found

142 | Measuring environmental variables Table 5.2 (continued) Category

Comments

Dew

Heavy morning dews can be a massive mortality factor for small insects and mites in exposed situations (Samways (1979) Wind meters are available for measuring current wind speed, which is only meaningful for behavioural studies, and for standardizing sampling, e.g. sampling is only done on windless days ; for ecological studies, it may be more important to have longterm records from a nearby weather station, or, where extent of exposure is required, it is necessary to install data loggers which monitor wind continuously; wind can interact with rainfall, and together, they can have storm impact

Wind

Cloud cover

Measurement of cloud cover is very important when sampling flying insects, and it is important to monitoring under standardized conditions, e.g.